Ultrasound technology to control the spatial organization of cells and proteins in engineered tissues

ABSTRACT

The present invention is directed to methods of inducing spatial organization of cells an in vitro culture system using ultrasound technology. The invention is further directed to methods of inducing extracellular matrix remodeling and neovessel formation in an in vitro culture system and generating vascularized engineered tissue constructs using ultrasound technology.

This application is a divisional of U.S. patent application Ser. No.13/321,218, filed Jan. 30, 2012, which is a national stage applicationunder 35 U.S.C. §371 of PCT Application No. PCT/US2010/031281, filedApr. 15, 2010, which claims the benefit of 61/179,646, filed May 19,2009, which are hereby incorporated by reference in their entirety.

This invention was made with government support under grant numberR01EB008368 awarded by the National Institutes of Health. The governmenthas certain rights in this invention.

FIELD OF THE INVENTION

The present invention is directed to methods of inducing cellularspatial organization, extracellular remodeling, and neovessel formationin an in vitro tissue culture system using ultrasound technology.

BACKGROUND OF THE INVENTION

Tissue and organ transplantation is a worldwide therapeutic approach forend-stage organ failure (Nasseri et al., “Tissue Engineering: AnEvolving 21st-Century Science to Provide Biological Replacement forReconstruction and Transplantation,” Surgery 130:781-784 (2001)).Currently, over 100,000 people are in need of an organ transplant in theUnited States alone. Each day, 17-20 patients die while waiting fordonor organs. These statistics highlight the worsening problem facingtransplant patients—the demand for tissues and organs far outweighs theavailable supply. The field of tissue engineering offers great potentialfor reducing the number of patient deaths associated with this shortageof organs. By developing methods for repairing or replacing diseased orinjured tissues and organs, tissue engineering aims to provide analternative supply of tissues and organs to balance supply and demand(Langer et al., “Tissue Engineering,” Science 260:920-926 (1993)).Before such a goal can be fully realized, tissue engineers need tosuccessfully reconstitute viable tissues and organs in vitro, a taskthat depends on the delivery of essential nutrients and oxygen to allcells within the tissue to uphold their metabolic processes. Existingdelivery methods include the passive diffusion of oxygen and nutrientsthrough the tissue and host-dependent vascularization of the tissueafter implantation (Nomi et al., “Principals of Neovascularization forTissue Engineering,” Molecular Aspects of Medicine 23:463-483 (2002);Lokmic et al., “Engineering the Microcirculation,” Tissue Engineering14(1):87-103 (2008); Tremblay et al., “Inosculation of Tissue-EngineeredCapillaries with the Host's Vasculature in a Reconstructed SkinTransplanted on Mice,” American Journal of Transplantation 5:1002-1010(2005)). These methods are limited to tissues with less than a fewmillimeters in thickness and have therefore, only been successfully usedfor the development of skin replacements (Folkman et al.,“Self-Regulation of Growth in Three Dimensions,” The Journal ofExperimental Medicine 138:745-753 (1973); Mooney et al., “Growing NewOrgans,” Scientific American 280(4):60-65 (1999); Nomi et al.,“Principals of Neovascularization for Tissue Engineering,” MolecularAspects of Medicine 23:463-483 (2002); Tremblay et al., “Inosculation ofTissue-Engineered Capillaries with the Host's Vasculature in aReconstructed Skin Transplanted on Mice,” American Journal ofTransplantation 5:1002-1010 (2005); Griffith et al., “Diffusion Limitsof an in Vitro Thick Prevascularized Tissue,” Tissue Engineering11(1-2):257-266 (2005)). As such, the engineering of larger, morecomplex, three-dimensional (3D) tissues and organs requires the in vitrodevelopment of a vascular system throughout the construct to adequatelyprovide oxygen and nutrients to all areas of the tissue (Griffith etal., “Tissue Engineering-Current Challenges and ExpandingOpportunities,” Science 295:1009-1014 (2002); Nerem R. M., “TissueEngineering: The Hope, the Hype, and the Future,” Tissue Engineering12:1143-1150 (2006); Jain et al., “Engineering Vascularized Tissue,”Nature Biotechnology 23(7):821-823 (2005); Levenburg et al.,“Engineering Vascularized Skeletal Muscle Tissue,” Nature Biotechnology23(7):879-884 (2005)).

Successful induction of neovessel network formation in tissue constructsdepends on the stimulation of endothelial cell functions critical toangiogenesis. Endothelial cell survival, growth, migration anddifferentiation are influenced by the spatial distribution ofendothelial cells and the organization of surrounding extracellularmatrix (“ECM”) (Korff et al., “Integration of Endothelial Cells inMulticellular Spheroids Prevents Apoptosis and Induces Differentiation,”The Journal of Cell Biology 143(5):1341-1352 (1998); Ino et al.,“Application of Magnetic Force-Based Cell Patterning for ControllingCell-Cell Interactions in Angiogenesis,” Biotechnology andBioengineering 102(3):882-890 (2009); Vailhe et al., “In Vitro Models ofVasculogenesis and Angiogenesis,” Laboratory Investigation 81(4):439-452(2001); Nehls et al., “The Configuration of Fibrin Clots DeterminesCapillary Morphogenesis and Endothelial Cell Migration,” MicrovascularResearch 51:347-364 (1996); Vailhe et al., “In Vitro Angiogenesis isModulated by the Mechanical Properties of Fibrin Gels and is Related toAlpha-v Beta-3 Integrin Localization,” In Vitro Cell. Dev. Biol.-Animal33:763-773 (1997); Ingber et al., “Mechanochemical Switching betweenGrowth and Differentiation During Fibroblast Growth Factor-StimulatedAngiogenesis In Vitro: Role of Extracellular Matrix,” The Journal ofCell Biology 109:317-330 (1989); Stephanou et al., “The Rigidity infibrin Gels as a Contributing Factor to the Dynamics of In VitroVascular Cord Formation,” Microvascular Research 73:182-190 (2007);Sieminski et al., “The Relative Magnitudes of Endothelial ForceGeneration and Matrix Stiffness Modulate Capillary Morphogenesis InVitro,” Experimental Cell Research 297:574-584 (2004; and Montesano etal., “In Vitro Rapid Organization of Endothelial Cells intoCapillary-Like Networks is Promoted by Collagen Matrices,” The Journalof Cell Biology 97:1648-1652 (1983), which are hereby incorporated byreference in their entirety). As such, control over both endothelialcell and ECM protein organization within 3D tissue constructs willaffect endothelial cell functions essential to angiogenesis.

Technologies currently in development to organize cells and proteinsinto complex patterns can be divided into two general categories. In thefirst approach, micropatterning of cell-adhesive contacts usingextracellular matrix proteins coated onto microfabricated stamps byphotolithography or microcontact printing is used to direct celladhesion into pre-designed patterns. In the second approach, a force isapplied to cells to direct cell movement to a desired location. Theapplied force can be optical, magnetic, electrokinetic, or fluidic ((Linet al., “Dielectrophoresis Based-Cell Patterning for TissueEngineering,” Biotechnol J 1:949-57 (2006)). The ability of acousticradiation forces associated with ultrasound standing wave fields tocontrol the spatial distribution of cells and extracellular matrixproteins in a three-dimensional tissue model has not previously beeninvestigated.

When an ultrasonic pressure wave is incident on an acoustic reflector,the reflected wave interferes with the incident wave resulting in thedevelopment of an ultrasound standing wave field (USWF). An USWF ischaracterized by areas of maximum pressure, known as pressure antinodes,and areas of zero pressure, known as pressure nodes. Exposure ofparticle or cell suspensions to an USWF can result in the alignment ofparticles or cells into bands that are perpendicular to the direction ofsound propagation and that are spaced at half-wavelength intervals(Coakley et al., “Cell Manipulation in Ultrasonic Standing Wave Fields,”J Chem Tech Biotechnol 44:43-62 (1989); Gould et al., “The Effects ofAcoustic Forces on Small Particles in Suspension,” In: Finite amplitudewave effects in fluids: Proceedings of the 1973 Symposium, Guildford:IPC Science and Technology Press Ltd, Bjorno L, ed. pp. 252-7 (1974);and Whitworth et al., “Particle Column Formation in a StationaryUltrasonic Field,” J Acoust Soc Am 91:79-85 (1992)). A primary acousticradiation force, (F_(rad)), generated along the direction of soundpropagation in the USWF, is largely responsible for this movement.F_(rad) is defined as

$\begin{matrix}{F_{rad} = {\left( \frac{{- \pi}\; P_{o}^{2}V\; \beta_{o}}{2\lambda} \right)*\varphi*{\sin \left( \frac{4\pi \; z}{\lambda} \right)}}} & \left( {{Equation}\mspace{14mu} 1} \right)\end{matrix}$

where P_(o) is the USWF peak pressure amplitude, V is the sphericalparticle volume, λ is the wavelength of the sound field, z is theperpendicular distance on axis from pressure nodal planes, and φ is anacoustic contrast factor given by

$\begin{matrix}{\varphi = {\frac{{5\rho_{p}} - {2\rho_{o}}}{{2\rho_{p}} + \rho_{o}} - \frac{\beta_{p}}{\beta_{o}}}} & \left( {{Equation}\mspace{14mu} 2} \right)\end{matrix}$

where ρ_(p) and β_(p) are the density and compressibility of theparticles or cells, and ρ_(o) and β_(o) are the density andcompressibility of the suspending medium, respectively (Gol'dberg Z A,“Radiation Forces Acting on a Particle in a Sound Field,” In: HighIntensity Ultrasonic Fields, New York: Plenum Press, Rozenberg L D, ed.,pp. 109-17 (1971); Gor'kov L P, “On the Forces Acting on a SmallParticle in an Acoustical Field in an Ideal Fluid,” Sov Phys Dokl6:773-5 (1962); and Gould et al., “The Effects of Acoustic Forces onSmall Particles in Suspension,” In: Finite amplitude wave effects influids: Proceedings of the 1973 Symposium, Guildford: IPC Science andTechnology Press Ltd, Bjorno L, ed. pp. 252-7 (1974). The forcesgenerating the banded pattern exist only during application of the USWF.Therefore, to maintain the USWF-induced banded distribution, suspendingmediums have been used that undergo a phase conversion from a liquid toa solid state during USWF exposure (Gherardini et al., “A Study of theSpatial Organisation of Microbial Cells in a Gel Matrix Subjected toTreatment With Ultrasound Standing Waves,” Bioseparation 10:153-62(2002); Gherardini et al., “A New Immobilisation Method to ArrangeParticles in a Gel Matrix by Ultrasound Standing Waves,” Ultrasound MedBiol 31:261-72 (2005); Saito et al., “Fabrication of a Polymer CompositeWith Periodic Structure by the Use of Ultrasonic Waves,” J Appl Phys83:3490-4 (1998); and Saito et al.,” “Composite Materials WithUltrasonically Induced Layer or Lattice Structure,” Jpn J Appl Phys38:3028-31 (1999)). In this way, the banded pattern is retained afterremoval of the sound field.

The present invention is directed to overcoming these and otherdeficiencies in the art.

SUMMARY OF THE INVENTION

A first aspect of the present invention is directed to a method ofinducing spatial organization of cells in an in vitro culture system.This method involves providing an in vitro culture system having cellsand a biological support material and placing the in vitro culturesystem in an ultrasound exposure chamber. The method further involvesexposing the in vitro culture system to an ultrasound standing wavefield under conditions effective to induce cellular spatial organizationand incubating the in vitro culture system containing the spatiallyorganized cells under conditions effective to permit cell behaviorimportant for tissue generation.

A second aspect of the present invention is directed to a method ofinducing neovessel formation in an in vitro culture system. This methodinvolves providing an in vitro culture system comprising a biologicalsupport material and endothelial cells, and placing the in vitro culturesystem in an ultrasound exposure chamber. The method further involvesexposing the in vitro culture system to an ultrasound standing wavefield under conditions effective to spatially organize endothelialcells, and incubating the in vitro culture system containing thespatially organized endothelial cells under conditions effective toinduce neovessel formation.

Another aspect of the present invention is directed to a vascularizedengineered tissue construct. This vascularized engineered tissueconstruct has a three-dimensional thickness of at least 2 mm.

Methods of promoting angiogenesis throughout three dimensionalengineered tissue are needed to fabricate large vascularized tissues andorgans for the field of tissue engineering. Angiogenic cell behaviorsare modulated by the spatial arrangement of endothelial cells, theorganization of their surrounding extracellular matrix (ECM), and bymechanical force application. The present invention demonstrates thatultrasound technology is capable of controlling all three of theseregulatory factors and can induce spatial organization of both cells andproteins and subsequently vascularization within three dimensionalengineered tissue constructs. The present invention will allow for thesuccessful reconstitution of viable tissues and organs in vitro, makingit possible to repair or replace diseased or injured tissues and organs.

BRIEF DESCRIPTION OF THE DRAWINGS

FIGS. 1A-1B are schematic representations of the experimental set-up forexposing in vitro cell culture systems to ultrasound standing wave field(UWSF). FIG. 1A shows the ultrasound wave field exposure system. A thickrubber absorber is placed above a sealed, submerged sample holder forultrasound traveling wave field (UTWF) exposures. FIG. 1B is an enlargedview of the silicone elastomer-bottomed (acoustic attenuation of 0.06dB) sample holder (FlexCell Inc., Hillsborough, N.C.). Well diameterswere decreased to 1 cm using Sylgard 184® Silicone Elastomer (DowCorning, Midland, Mich.) molds to fit the sample size to the dimensionsof the ultrasound beam.

FIGS. 2A-2D are graphs showing the ultrasound beam patterns in freefield and sample space. Transaxial spatial distributions in pressurewere measured in both the free field (FIG. 2A) and within the 1 cmdiameter sample space (FIG. 2B). Data are plotted as normalized pressureversus spatial location. The calculated −3 dB and −6 dB beam widths were0.8 cm and 1.2 cm in the free field. The axial spatial distribution inpressure within an USWF (FIGS. 2C-2D) was measured in 0.2 mm intervalsthrough a 1 cm distance below the air interface in both the free field(FIG. 2C) and sample space (FIG. 2D). Peak positive pressures weremeasured for each position. The distance between pressure nulls wascalculated as the distance between pressure minima and was approximatelythe expected half-wavelength spacing for 1 MHz sound in water (0.75 mm).

FIGS. 3A-3C illustrate ultrasound standing wave fields (USWF) inducedchanges in the spatial organization of cells in 3D collagen gels.Fibronectin-null (FN−/−) myofibroblasts (MF) at 2×10⁵ c/ml (FIGS. 3A and3B) or 4×10⁶ c/ml (FIG. 3C), suspended in a collagen I solution (0.8mg/ml), were exposed to an USWF using a 1 MHz continuous wave (CW)sinusoidal signal for 15 min with an incident pressure amplitude of 0.1MPa. Representative phase-contrast images, from one of at least twoexperiments performed in duplicate, show dark bands of cells inUSWF-exposed gels (FIGS. 3B and 3C) that are absent in sham gels (FIG.3A) where a homogeneous cell distribution is observed. Scale bar, 200μm.

FIG. 4 is a graph showing that USWF exposure does not affect cellviability. Fibronectin-null myofibroblasts (2×10⁵ c/ml) suspended in acollagen I solution (0.8 mg/ml) were exposed to an USWF using a 1 MHz CWsinusoidal signal for 15 min with an incident pressure amplitude of 0.1MPa. Following a 20 hr incubation period at 37° C. and 8% CO₂,cell-embedded collagen gels were incubated with MTT for 4 hrs, digestedwith collagenase, and formazen crystals were dissolved in acidifiedisopropanol to measure the absorbance at 570 nm and 700 nm (background)for quantification of viable cell number. Data from four experimentswere normalized to sham average absorbance values±SEM.

FIGS. 5A-5B are photomicrographs showing the spatial localization ofsoluble fibronectin (FN) in 3D collagen gels with and without USWFexposure. Fluorescently labeled fibronectin (FN-488; 10 μg/ml) was addedto collagen I solutions in the absence of cells, and exposed to an USWFusing a 1 MHz CW sinusoidal signal for 15 min with an incident pressureamplitude of 0.1 MPa (FIG. 5A, bottom panels). Fibronectin distributionwas analyzed using phase-contrast (FIG. 5A; left panel) and fluorescentmicroscopy (FIG. 5A; right panel). Fibronectin-null myofibroblasts insuspension were incubated with 100 μg/ml FN-488 in the presence of 1 mMMnCl₂. Cells were washed twice to remove unbound FN and were then addedto unpolymerized collagen I solutions (4×10⁶ c/ml) for exposure to theaforementioned USWF. Cell and FN distribution were analyzed usingphase-contrast (FIG. 5B, left panel) and fluorescent microscopy (FIG.5B, right panel), respectively. Representative images are shown. Scalebar, 200 μm

FIGS. 6A-6D show USWF induced cellular organization enhancescell-mediated collagen gel contraction. Fibronectin-null myofibroblasts(2×10⁵ c/ml) were suspended in a collagen I solution (FIGS. 6A and 6B)or embedded in polymerized collagen I gels (FIGS. 6C and 6D) and wereexposed to an USWF using a 1 MHz CW sinusoidal signal for 15 min with anincident pressure amplitude of 0.1 MPa. Following USWF exposure,floating gels were incubated for 20 hrs at 37° C. and 8% CO₂ at whichtime they were removed from the wells and weighed. Percent contraction(FIGS. 6A and 6C) was calculated as (1−(test gel weight/no-cell gelweight))×100. Data are average values from four experiments performed inquadruplicate. *p<0.05 vs. sham by paired t-test. Cell distribution(FIGS. 6B and 6D) in USWF-exposed (right) and sham gels (left) wasanalyzed using phase-contrast microscopy to correlate changes in percentcontraction with USWF-mediated changes in the spatial distribution ofcells. Scale bar, 200 μm.

FIG. 7 is a series of photomicrographs illustrating cell banding as afunction of USWF pressure amplitude. Fibronectin-null cells (4×10⁶cell/ml) were suspended in collagen I solutions and were exposed at roomtemperature for 15 min to a continuous wave USWF (1 MHz source) withvarious peak pressure amplitudes. Representative phase-contrast images,from one of four experiments, indicate cell banding in samples exposedto 0.1 MPa and above. Scale bar, 200 μm.

FIGS. 8A-8B are graphs showing the biphasic effect of USWF pressureamplitude on cell-mediated collagen gel contraction. In FIG. 8A,fibronectin-null cells (4×10⁶ cell/ml) suspended in collagen I solutionswere exposed at room temperature for 15 min to a continuous wave USWF (1MHz source) with various peak pressure amplitudes. Following 1 hrincubation at 37° C. and 8% CO₂, gel diameters were measured. Data arepresented as average percent radial gel contraction±SEM and arenormalized to the sham condition (n=10). The * indicates a differencefrom sham group (p<0.05). In FIG. 8B, fibronectin-null cells (2×10⁵cell/ml) suspended in a collagen I solutions were exposed at roomtemperature for 15 min to a continuous wave USWF (1 MHz source) withpeak pressure amplitude of 0.3 MPa. Following 1 hr incubation at 37° C.and 8% CO₂, cell viability was assessed using MTT. Data are presented asaverage fold difference in absorbance±SEM normalized to sham averageabsorbance values (n=3; p>0.05).

FIG. 9 is a series of fluorescent photomicrographs showing the biphasiceffect of USWF pressure amplitude on cell-mediated collagenreorganization. Fibronectin-null cells (4×10⁶ cell/ml) suspended incollagen I solutions were exposed at room temperature for 15 min to acontinuous wave USWF (1 MHz source) with various peak pressureamplitudes (0-0.30 MPa). Following 1 hr incubation at 37° C. and 8% CO₂,gels were fixed with 4% paraformaldehyde and imaged usingsecond-harmonic generation microscopy as described infra. Representativemerged images, from one of three experiments, are shown. Scale bar, 100μm.

FIGS. 10A-10B shows endothelial cell sprouts emerging from USWF-inducedendothelial cell bands. Human umbilical vein endothelial cells (HUVEC;1×10⁶ c/ml) suspended in a collagen I solution were exposed to an USWFusing a 1 MHz CW sinusoidal signal for 15 min with an incident pressureamplitude of 0.1 MPa. Following USWF exposure, gels were incubated for24 hrs at 37° C. and 5% CO2 and then imaged using phase-contrastmicroscopy. Endothelial cell sprouts (arrows) can only be seen inUSWF-exposed gels (FIG. 10B) and not in sham gels (FIG. 10A). Imagesrepresent similar results obtained from six experiments performed intriplicate. Scale bar, 100 μm.

FIGS. 11A-11B are a series of photomicrographs tracking thecapillary-like endothelial cell sprouts emerging from USWF-inducedendothelial cell bands. HUVEC were suspended at 1×10⁶ cell/ml in aneutralized type-I collagen solution and were exposed to a 1 MHz USWFwith a peak pressure amplitude of 0.2 MPa for a 15 min duration at roomtemperature to promote the formation of multicellular endothelial cellbands (FIG. 11B). Sham samples were treated in the exact same manner asUSWF-exposed samples but did not receive USWF treatment (FIG. 11A).Representative phase-contrast images collected at the indicated timepoints following USWF exposure are shown. Multiple capillary-likesprouts can be seen emerging from the endothelial cell banded area.Arrow, same sprout over time. Scale bar, 100 μm.

FIGS. 12A-12B are fluorescent photomicrographs showing that USWF-inducedendothelial cell sprouts are multicellular structures. Four daysfollowing Sham (FIG. 12A) or USWF (FIG. 12B) exposure, samples werefixed in 4% paraformaldehyde and permeabilized with 0.5% TritonX-100.Cell nuclei were visualized by staining with DAPI and HUVEC werevisualized by staining with anti-human CD31 monoclonal antibody followedby AlexaFluor-594 conjugated anti-mouse IgG. Two-photon microscopy wasused to collect images along the z-axis in 1 μm slices. Images were thenprojected onto the z-plane using ImageJ software. Scale bar, 15 μm.

FIGS. 13A-13B are photomicrographs demonstrating that USWF-inducedendothelial cell sprouts contain endothelial cell-lined lumen. Four daysfollowing USWF exposure (FIG. 13B), samples were fixed in 4%paraformaldehyde and processed normally for histology. Sham samples weretreated in the exact same manner as USWF-exposed samples but did notreceive USWF treatment (FIG. 13A). Four-micrometer thick gelcross-sections were stained with H&E to differentiate cells from thesurrounding collagen matrix. HUVEC-lined lumens in representative imagesare indicated by the arrows. Scale bar, 100 μm.

FIGS. 14A-14B are fluorescent photomicrographs showing reorganization ofcollagen fibers surrounding USWF-induced capillary sprouts in thedirection of sprout outgrowth. One day following USWF exposure, sampleswere fixed in 4% paraformaldehyde (FIG. 14B). Sham samples were treatedin the exact same manner as USWF-exposed samples but did not receiveUSWF treatment (FIG. 14A). Collagen type-I fibers were visualized usingsecond harmonic generation microscopy imaging on a two-photonmicroscope. HUVEC were visualized using their intrinsicauto-fluorescence. Scale bar, 1 μm.

DETAILED DESCRIPTION OF THE INVENTION

A first aspect of the present invention is directed to a method ofinducing spatial organization of cells in an in vitro culture system.This method involves providing an in vitro culture system having cellsand a biological support material and placing the in vitro culturesystem in an ultrasound exposure chamber. The method further involvesexposing the in vitro culture system to an ultrasound standing wavefield under conditions effective to induce cellular spatialorganization, and incubating the in vitro culture system containing thespatially organized cells under conditions effective to permit cellbehavior important for tissue generation.

As used herein, an in vitro culture system refers to any two or threedimensional culture of living cells or tissue, preferably mammaliancells or tissue, produced primarily by growth in vitro. The in vitroculture system of the present invention may include one or more types ofcells or tissues. The cells of the in vitro system may be primary cellcultures, cell lines, or a combination of both. Suitable cell typesinclude, but are not limited to, smooth muscle cells, cardiac musclecells, cardiac myocytes, platelets, epithelial cells, endothelial cells,endothelial progenitor cells, urothelial cells, fibroblasts, embryonicfibroblasts, myoblasts, chondrocytes, chondroblasts, osteoblasts,osteoclasts, keratinocytes, hepatocytes, bile duct cells, pancreaticislet cells, thyroid, parathyroid, adrenal, hypothalamic, pituitary,ovarian, testicular, salivary gland cells, adipocytes, embryonic stemcells, mesenchymal stem cells, hematopoietic cells, neural cells, andprecursor cells.

In accordance with this aspect of the present invention, the in vitroculture system containing spatially organized cells is incubated underconditions effective to permit cell behavior important for tissuegeneration. Tissue generation as used herein refers to both de novotissue generation and tissue regeneration. Cell behaviors that areinvolved in tissue generation include, without limitation, cellsurvival, cell growth, cell differentiation, cell migration, changes ingene expression, and extracellular matrix remodeling. Tissue generationcan be measure or assessed by any one or more of the above identifiedcell behaviors.

In one embodiment of the present invention, the in vitro tissue culturesystem is one having a configuration that is amenable to the generationof an engineered tissue construct. An engineered tissue construct is athree dimensional mass of living mammalian tissue produced primarily bygrowth in vitro that shares critical structural and functionalcharacteristics with intact tissue, such as distinctive multicellularorganization and oriented contractile function. Engineered tissueconstructs generated using the methods of the present invention can beany desired tissue construct including, but not limited to, a muscularconstruct, a vascular construct, an esophageal construct, an intestinalconstruct, a rectal construct, an ureteral construct, a cartilaginousconstruct, a cardiac construct, a liver construct, a bladder construct,a kidney construct, a pancreatic construct, a skeletal construct, afilamentous/ligament construct, a lung construct, a neural construct, abone construct, and a skin construct.

In accordance with this aspect of the present invention, the biologicalsupport material of the in vitro culture system consists of a biologicalmaterial that supports the growth and survival of living cells andtissue under in vitro conditions. In an embodiment of the presentinvention, the biological support material of the in vitro culturesystem is a polymerizable culture media. Suitable polymerizable culturemedia solutions include, without limitation, collagen, collagen type-I,alginate, growth factor reduced Matrigel, Matrigel, hydrogel, andfibrin, any of which may be supplemented with suitable proteinaceousmaterials (e.g., chitosan, hyaluronan, polyethylene oxide/polypropyleneoxide). The gel may further include one or more of laminin, fibrin,fibronectin, proteoglycans, glycoproteins, glycosaminoglycans,chemotactic agents, or growth factors, for example, cytokines,eicosanoids, or differentiation factors. Upon polymerization of theculture media solution, a three-dimensional culture environment isformed.

In another embodiment of the present invention, the in vitro culturesystem includes a biological support material that promotes threedimensional cellular and tissue growth and expansion. In general, thesesupports are three-dimensional and are processable to form scaffolds ofa desired shape for the tissue of interest. Suitable three dimensionalbiological supports include, without limitation, filaments, meshes,foams, gels, ceramics, and acellularized extra-cellular matrix material.The biological support substrate preferably consists of a biocompatiblematerial, e.g., a biocompatible polymer having properties orincorporating modifications conducive to cell adherence and/or growth.In one embodiment, the support material is a porous polymer as describedin U.S. Pat. No. 6,103,255 to Levene, which is hereby incorporated byreference in its entirety. In another embodiment, the support materialis biodegradable or bioerodable, including those materials thathydrolyze slowly under physiological conditions. Suitable materials,include synthetic polymeric materials such as polyesters,polyorthoesters, polylactic acid, polyglycolic acid, polycaprolactone,or polyanhydrides, including polymers or copolymers of glycolic acid,lactic acid, or sebacic acid. Substrates comprising proteinaceouspolymers are also suitable for use in the methods of the presentinvention. Collagen gels, collagen sponges and meshes, and substratesbased on elastin, fibronectin, laminin, or other extracellular matrix orfibrillar proteins may also be employed. Either synthetic polymers orproteinaceous polymers may be modified or derivatized in any of avariety of ways, e.g., to increase their hydrophilicity and/or provideimproved cell adhesion characteristics. In certain embodiments of thepresent invention, the substrate may be coated with an agent, e.g.,denatured collagen, prior to seeding in order to increase cellularadherence. Materials useful as substrates for growing cells to producetissue engineered substrates, and methods of producing such substratesare known in the art and are described in U.S. Pat. No. 5,770,417 toVacanti et al., which is hereby incorporated by reference in itsentirety.

Other suitable biological support materials that can be used in themethods of the present invention include the multilayer scaffolddescribed in U.S. Pat. No. 6,143,292 to Weiss et al., which is herebyincorporated by reference in its entirety; the three dimensionalgeometric biocompatible porous scaffold described in U.S. Pat. No.6,206,924 to Timm, which is hereby incorporated by reference in itsentirety; the three dimensional matrix containing fibrin matrixdescribed in U.S. Patent Application Publication No. 20030166274; andthe hyaluronan based biodegradable scaffold described in U.S. Pat. No.5,939,323, which is hereby incorporated by reference in its entirety.

The in vitro culture system is cultured or maintained using standardtissue culture procedures. Appropriate growth and culture conditions forvarious mammalian cell types are well known in the art. The cells in thein vitro cell culture system may be seeded onto and/or within asubstrate from a suspension so that they are evenly distributed at arelatively high surface and/or volume density. The cell suspensions maycomprise approximately about 1×10⁴ to about 5×10⁷ cells/ml of culturemedium, or approximately about 2×10⁶ cells/ml to about 2×10⁷ cells/ml,or approximately about 5×10⁶ cells/ml. The optimal concentration andabsolute number of cells will vary with cell type, growth rate of thecells, substrate material, and a variety of other parameters. Thesuspension may be formed in any physiologically acceptable medium,preferably one that does not damage the cells or impair their ability toadhere to the substrate. Appropriate mediums include standard cellgrowth media (e.g., DMEM with 10% FBS).

Examples of suitable seeding and culturing methods for the growth ofthree-dimensional cell cultures, including techniques for establishing athree-dimensional matrix, inoculating the matrix with the desired cells,and maintaining the culture are disclosed in U.S. Pat. No. 6,537,567 toNiklason et al., U.S. Pat. No. 5,266,480 to Naughton et al., and U.S.Pat. No. 5,770,417 to Vacanti et al., which are hereby incorporated byreference in their entirety. Alternatively, the cells of the in vitrosystem can be suspended in a polymerizable cell media and seeded into anappropriate culture dish or onto an appropriate substrate as describedherein. Suitable substrates can be flat, tubular, or configured toassume any desired three-dimensional shape (e.g., spheres, ellipsoids,disks, sheets, or films as well as hollow spheres, hollow ellipsoids,and open-ended, hollow tubes).

Cells of the in vitro culture system are cultured in a media thatgenerally includes essential nutrients and, optionally, additionalelements such as growth factors, salts, minerals, vitamins, etc., thatmay be selected according to the cell type(s) being cultured. A standardgrowth media includes Dulbecco's Modified Eagle Medium, low glucose(DMEM), with 110 mg/L pyruvate and glutamine, supplemented with 10-20%fetal bovine serum (FBS) or calf serum and 100 U/ml penicillin. Theculture media may also contain particular growth factors selected toenhance cell survival, differentiation, secretion of specific proteins,etc. In accordance with this aspect of the present invention, factorsthat enhance cell growth, proliferation, and differentiation may beadded to the in vitro culture system.

In accordance with this aspect of the present invention the in vitroculture system may further contain one or more particles that are alsoresponsive to the ultrasound wave field exposure as described herein.These particles include, without limitation, nanoparticles,microparticles, microbubbles, and cells. In a preferred embodiment ofthe present invention, these particles contain or have bound theretobiologically active peptides, proteins, or peptide or protein mimetics.Suitable peptides, proteins, and protein mimetics include for examplegrowth factors (e.g., VEGF or FGF), adhesion proteins, extracellularmatrix proteins (e.g., fibronectin, recombinant fibronectin fragments,or vitronectin), angiogenic factors, etc. In a preferred embodiment ofthe present invention, the biologically active peptide, protein, orprotein mimetic is a biologically active fibronectin peptide, protein,or protein mimetic.

Once the desired in vitro culture system has been established, theculture system is exposed to an ultrasound wave field under conditionseffective for inducing cellular spatial organization. This spatialorganization of cells within the in vitro culture system facilitates keycellular functions, such as survival, growth, migration, anddifferentiation. In addition, as demonstrated herein, the spatialorganization of cells in the in vitro culture system facilitatesextracellular matrix remodeling thereby enhancing the mechanicalstrength of the cultured cells and tissue.

Ultrasound (“US”) is a form of mechanical energy that travels through amedium of propagation as an acoustic pressure wave at frequencies above20 kHz. It is used widely in the medical field as both a diagnostic andtherapeutic tool (Duck et al., “Ultrasound In Medicine,” Philadelphia:Institute of Physics Publishing (1998), which is hereby incorporated byreference in its entirety). US can interact with biological tissuesthrough thermal or mechanical mechanisms (Dalecki D., “MechanicalBioeffects of Ultrasound,” Annu. Rev. Biomed. Eng. 6:229-248 (2004),which is hereby incorporated by reference in its entirety). Mechanicalinteractions, associated with the generation of US-induced mechanicalforces in the medium of propagation, can produce US bioeffects (DaleckiD., “Mechanical Bioeffects of Ultrasound,” Annu. Rev. Biomed. Eng.6:229-248 (2004), which is hereby incorporated by reference in itsentirety). For example, red blood cells redistribute to equally spacedintervals due to USWF radiation forces (Dyson et al., “The Production ofBlood Cell Stasis and Endothelial Damage in the Blood Vessels of ChickEmbryos Treated with Ultrasound in a Stationary Wave Field,” Ultrasoundin Medicine and Biology 1:133-148 (1974), which is hereby incorporatedby reference in its entirety). Also, low-intensity pulsed US enhancesnitric oxide production in endothelial cells, a well-documentedendothelial cell response to shear stress and stimulates fibroblastgrowth through a signaling pathway known to be triggered by directmechanical stimulation (Altland et al., “Low-Intensity UltrasoundIncreases Endothelial Cell Nitric Oxide Synthase Activity and NitricOxide Synthesis,” Journal of Thrombosis and Haemostasis 2:637-643 (2004)and Zhou et al., “Molecular Mechanisms of Low Intensity PulsedUltrasound in Human Skin Fibroblasts,” The Journal of BiologicalChemistry 279(52):54463-54469 (2004), which are hereby incorporated byreference in their entirety).

In accordance with the methods of the present invention, ultrasoundexposure mediated “spatial organization” of cells in the in vitro cellculture system encompasses the alignment of cells into parallel sheets,planes, columns, and/or grid matrices. As demonstrated herein, exposureof the in vitro culture system of the present invention to an ultrasoundstanding wave field results in the alignment of cells in the system intobands that are perpendicular to the direction of sound propagation andthat are spaced at half-wavelength intervals. As discussed supra, the invitro cultures system of the present invention may also include one ormore particles that are also responsive to ultrasound exposure mediatedspatial organization. In one embodiment of the present invention,exposure of the particles (e.g. proteins bound to cells) to anultrasound wave field will localize the particles to the pressure node,thereby co-localizing the particles with the cells of the in vitroculture system. Co-localization of the particles and cells will providecells of the system the necessary and specific signals and growthfactors that mediate cell growth, proliferation, or differentiation in acontrolled manner. In another embodiment of the present invention,exposure of the particles (e.g. microbubbles) to an ultrasound wavefield will localize the particles to the pressure anti-nodes resultingin a staggered arrangement of the particles and cells of the in vitroculture system. This staggered arrangement of particles and cells willgenerate a concentration gradient of growth factors or matrix proteinscontained in the particles between the spatially organized cells. Theconcentration gradients will enhance phenotypic differentiation of thespatially organized cells (e.g., enhance neovessel formation ofendothelial cell bands).

Prior to ultrasound wave field exposure, the in vitro culture system isplaced in a suitable ultrasound exposure chamber, such as the onedescribed herein. This exposure chamber consists of a degassed chamberof deionized water in a plastic exposure tank. The in vitro culturesystem is sealed and submerged into the exposure tank via a sampleholder. The sample holder aligns the in vitro culture system with theultrasound wave beam to facilitate direct exposure. The exposure chambercan further include a means for controlling the sample/air interface(e.g., a rubber absorber) to allow for ultrasound standing or travelingwave fields to be generated in the same exposure chamber.

The parameters of ultrasound wave field exposure of the in vitro culturesystem will vary depending on the cell type, biological supportmaterial, and desired endpoint (e.g. spatial organization), and shouldbe optimized for each in vitro culture system to produce optimalresults. These parameters include, for example, acoustic pressureamplitude, frequency, and exposure duration. Methods for optimizingthese ultrasound exposure parameters are described herein.

In accordance with this aspect of the present invention, the in vitroculture system is exposed to an ultrasound standing wave field. Thecells and particles of the in vitro culture system may be exposed to oneor more ultrasound standing wave beams at one time depending on thedesired spatial organization. Exposure to one beam of ultrasound issufficient for spatially organizing cells in planes and columns.Alternatively, multiple intersecting beams of ultrasound can be used tospatially arrange cells and/or particles into a grid matrix and columns.Various transducer geometries and set-ups can be utilized to achieve thedesired spatial arrangement or organization of cells in the system. Theultrasound standing wave field exposure may involve exposure of the invitro culture system to a continuous wave signal or a pulsating wavesignal. If a pulsating wave signal exposure is employed the appropriatepulse frequency and duration must be optimized.

In accordance with this aspect of the present invention, exposure to theultrasound standing wave field is carried out at a pressure amplitudethat is optimal for mediating the spatial organization of cells in thein vitro culture system. In one embodiment, cells of the in vitroculture system are exposed to ultrasound pressure amplitude of about0.01 MPa to about 0.5 MPa. Likewise, the ultrasound frequency should bealso be optimized to ensure a frequency that promotes spatialorganization and neovessel formation is employed. In one embodiment, theexposure of the in vitro culture system to an ultrasound energy sourceis at a frequency of from about 0.02 MHz to about 20 MHz. Morepreferably the exposure to an ultrasound energy source is at a frequencyof from about 0.1 MHz to about 3 MHz. Most preferably, the ultrasound isapplied at a frequency of about 1 MHz.

The duration of ultrasound exposure will vary depending on the tissuetype of the in vitro culture system. In one embodiment, ultrasoundexposure is carried out for a period of about 10 seconds to about 60minutes. In another embodiment the exposure is for a period of about 60seconds to about 20 minutes. In another embodiment, the ultrasound isapplied for about 15 minutes. Exposure of the in vitro culture system tothe ultrasound standing wave may be repeated one or more times a day, aweek, or a month.

In another embodiment of the present invention, the spatially organizedcells of the in vitro culture system are further exposed to anultrasound traveling wave field. Exposure to an ultrasound travelingwave field will facilitate cell growth, survival, proliferation, andphenotypic differentiation of the spatially organized cells of the invitro culture system. Exposure of the spatially organized cells of thein vitro system can be carried out for any suitable duration of time.Preferably, the duration of exposure is between about 10 seconds andabout 60 minutes. In another embodiment, the exposure is for a period ofabout 60 seconds to 20 minutes. In yet another embodiment, the exposureis carried out for a period of about 10 minutes. Exposure of the invitro culture system to the ultrasound traveling wave may be repeatedone or more times a day, a week, or a month.

The parameters of ultrasound traveling wave exposure must be optimizedin accordance with the particular in vitro culture system utilized basedon cell type, biological support material, and particle type. In oneembodiment, the exposure of the in vitro culture system to an ultrasoundenergy source is at a frequency of about 0.02 MHz to about 20 MHz. Inanother embodiment, the exposure to an ultrasound energy source is at afrequency of about 0.1 MHz to about 3 MHz. Alternatively, the ultrasoundis applied at a frequency of about 1 MHz.

Exposure of the in vitro culture system may consist of a continuoustraveling wave exposure or a pulsating traveling wave exposure. When apulsed ultrasound exposure is employed, the pulse frequency and durationmust also be optimized. In a preferred embodiment, the in vitro culturesystem is exposed to an ultrasonic field pulsed at 1 kHz with a 200 μspulse duration.

Therapeutic ultrasound treatments that enhance wound healing have beenhypothesized to do so by promoting the growth of blood vessels withinthe wound space (Young et al., “The Effect of Therapeutic Ultrasound onAngiogenesis,” Ultrasound in Medicine and Biology 16:261-269 (1990),which is hereby incorporated by reference in its entirety). In fact,similar ultrasound exposure protocols have been shown to enhance bothtissue healing and angiogenesis (Young et al., “The Effect ofTherapeutic Ultrasound on Angiogenesis,” Ultrasound in Medicine andBiology 16:261-269 (1990) and Dyson et al., “The Stimulation of TissueRegeneration by Means of Ultrasound,” Clinical Science 35:273-285(1968), which are hereby incorporated by reference in their entirety).Exposure to the aforementioned growth-promoting UTWF is expected toenhance neovessel formation in 3D collagen gels. Therefore, it may bedesirable to expose USWF-induced spatially organized cells to the UTWFexposure regimen as described supra. As therapeutic ultrasound exposureenhances both the number of blood vessels in the wound space (Barzelaiet al., “Low-Intensity Ultrasound Induces Angiogenesis in Rat Hind-LimbIschemia,” Ultrasound in Medicine and Biology 32(1):39-145 (2006) andYoung et al., “The Effect of Therapeutic Ultrasound on Angiogenesis,”Ultrasound in Medicine and Biology 16:261-269 (1990), which are herebyincorporated by reference in their entirety) and the length of newlyformed capillary structures (Mizrahi et al., “Ultrasound-InducedAngiogenic Response in Endothelial Cells,” Ultrasound in Medicine andBiology 33(11):1818-1829 (2007), which is hereby incorporated byreference in its entirety), it is expected that USWF-induced cell andprotein organization and UTWF exposure will combine to promoteangiogenesis for the vascularization of 3D tissue constructs.

Following ultrasound wave field exposure of the in vitro culture systemof the present invention, the culture system is maintained underconditions effective to facilitate cell or tissue survival, growth,proliferation, differentiation, gene expression, migration, andextracellular matrix remodeling. Suitable conditions will depend on thecells or tissue of the in vitro culture system.

A second aspect of the present invention is directed to a method ofinducing extracellular matrix remodeling in an in vitro culture system.This method involves providing an in vitro culture system having abiological support material and placing the in vitro culture system inan ultrasound exposure chamber. The method further involves exposing thein vitro culture system to an ultrasound standing wave field underconditions effective to induce extracellular matrix remodeling, andincubating the in vitro culture system under conditions to permitextracellular matrix remodeling.

As demonstrated in the Examples described herein, ultrasound exposure iscapable of mediating the spatial organization of cells and cell boundproteins within a three dimensional collagen gel. Accordingly,ultrasound technology provides a mean for regulating the movement ofextracellular matrix protein movement within an in vitro culture system.Spatial organization of extracellular matrix proteins, like fibronectin,in a collagen matrix enhances the extent of cell-mediated collagenremodeling. Controlling extracellular matrix remodeling within an invitro culture system is important for promoting and enhancing themechanical strength of the cultured tissue, a factor that is criticalfor engineering larger, more complex, three dimensional tissues than arecurrently available.

Any of the in vitro culture systems and the ultrasound exposureparameters described infra are suitable for use in accordance with thisaspect of the present invention.

A third aspect of the present invention is directed to a method ofinducing neovessel formation in an in vitro culture system. This methodinvolves providing an in vitro culture system comprising a biologicalsupport material and endothelial cells, and placing the in vitro culturesystem in an ultrasound exposure chamber. The method further involvesexposing the in vitro culture system to an ultrasound standing wavefield under conditions effective to spatially organize endothelialcells, and incubating the in vitro culture system containing thespatially organized endothelial cells under conditions effective toinduce neovessel formation.

In accordance with this aspect of the present invention, “neovesselformation” refers to the generation of any vascular structure includingcapillaries and blood vessels in the in vitro culture system. Inaccordance with this aspect of the invention, neovessel formation mayresult from neovascularization or angiogenic processes.

Accordingly, in a preferred embodiment of the present invention, the invitro culture system includes vascular endothelial cells in combinationwith one or more tissue specific cell types. Suitable endothelial cellsinclude primary endothelial cells (e.g., human umbilical veinendothelial cells) as well as endothelial cell lines. Also suitable foruse in the present invention are endothelial progenitor cells,hematopoietic cells, and embryonic stem cells capable of endothelialcell differentiation. Preferably, the endothelial cells and tissuespecific cells of the present invention are mammalian cells and morepreferably the cells are human cells.

The in vitro culture system may further contain particles (e.g.,nanoparticles, microparticles, microbubbles, etc.) that contain or carryendothelial cell specific growth factors and/or angiogenic growthfactors to promote endothelial cell differentiation and neovesselformation. Suitable growth factors include, without limitation, FGF,bFGF, acid FGF (aFGF), FGF-2, FGF-4, EGF, PDGF, TGF-betal,angiopoietin-1, angiopoietin-2, placental growth factor (PlGF), VEGF,PMA (phorbol 12-myristate 13-acetate), and the like.

The methods of the present invention can be adapted to any in vitroculture system known in the art. In fact, various in vitro culturesystems have been developed for the generation of three dimensionalengineered tissue construct and all are suitable for use in the methodof the present invention. It is expected that application of the methodsof the present invention to these systems will facilitate thevascularization of such tissue constructs resulting in the generation ofmore complex three dimensional tissues having expanded in vitro and,more importantly, having in vivo utility. Examples of suitable threedimensional tissue engineered constructs include, without limitation,oral tissue constructs (U.S. Patent Application Publication No.20060171902 to Atala et al., which is hereby incorporated by referencein its entirety); cardiac constructs (U.S. Patent ApplicationPublication No. 20080075750 to Akins and U.S. Pat. No. 5,885,829 toMooney et al, which are hereby incorporated by reference in theirentirety); embryonic brain tissue construct (U.S. Patent ApplicationPublication No. 20060030043 to Ma, which is hereby incorporated byreference in its entirety); muscular construct (U.S. Patent ApplicationPublication Nos. 2006019827 to Levenberg et al., 20060134076 to Bitar etal, and U.S. Pat. No. 6,537,567 to Niklason et al., which are herebyincorporated by reference in their entirety); stromal cell constructs(U.S. Pat. No. 4,963,489 to Naughton et al. and U.S. Patent ApplicationPublication No. 2003007954 to Naughton et al., which are herebyincorporated by reference in their entirety); embryonic tissueconstructs (U.S. Patent Application Publication No. 20050031598 toLevenberg et al., which is hereby incorporated by reference in itsentirety); pancreatic constructs (U.S. Pat. No. 6,022,743 to Naughton etal., which is hereby incorporated by reference in its entirety); skinconstructs (U.S. Pat. No. 5,266,480 to Naughton et al., which is herebyincorporated by reference in its entirety); filamentous tissue/ligamentconstruct (U.S. Pat. No. 6,140,039 to Naughton et al., U.S. Pat. No.6,840,962 to Vacanti et al., and now U.S. Pat. No. 6,737,053 to Goh etal., which are hereby incorporated by reference in their entirety);cartilage constructs (U.S. Pat. No. 5,902,741 to Purchio et al., whichis hereby incorporated by reference in its entirety); vascularconstructs (U.S. Pat. No. 6,455,311 to Vacanti, U.S. Pat. No. 7,112,218to McAllister et al., and U.S. Pat. No. 7,179,287 to Wolfinbarger, whichare hereby incorporated by reference in their entirety); kidneyconstructs (U.S. Pat. No. 5,516,680 to Naughton et al., which is herebyincorporated by reference in its entirety); uterine constructs (U.S.Patent Application Publication No. 20030096406 to Atala et al., which ishereby incorporated by reference in its entirety); and liver constructs(U.S. Pat. No. 5,624,840 to Naughton et al., which is herebyincorporated by reference in its entirety).

Any of the in vitro culture systems and the ultrasound exposureparameters described infra are suitable for use in accordance with thisaspect of the present invention.

Another aspect of the present invention is directed to a vascularizedengineered tissue construct made in accordance with the methodsdescribed herein. This vascularized engineered tissue construct has athree-dimensional thickness of at least 2 mm.

In accordance with this aspect of the present invention, thevascularized engineered tissue construct contains networks ofmicrovessels having internal diameters of about 0.5 mm or less. Thesemicrovessels of the tissue construct form an “exchange” network that iscapable of supplying nutrients and removing wastes from the new tissue.Upon transplantation of the vascularized tissue construct of the presentinvention, the tissue construct integrates or connects to host tissueeither by inducing incoming vessels into the construct (i.e.,angioinduction) and/or by allowing the construct vessels to meet thehost vessles (i.e., inosculation).

Vascularized engineered tissue constructs generated in accordance withthe present invention include, without limitation, a vascularizedcardiac construct, muscular construct, a vascular construct, anesophageal construct, an intestinal construct, a rectal construct, anureteral construct, a cartilaginous construct, a liver construct, abladder construct, a kidney construct, a pancreatic construct, askeletal construct, a filamentous/ligament construct, a lung construct,a neural construct, a bone construct, and a skin construct.

In one embodiment of the present invention, a three dimensionalvascularized engineered cardiac tissue construct is generated. The celltypes that may be used to generate a three dimensional cardiac constructmay include, but are not limited to, cardiomyocytes, endocardial cells,cardiac adrenergic cells, cardiac fibroblasts, vascular endothelialcells, smooth muscle cells, stem cells, cardiac progenitor cells, andmyocardial precursor cells. Depending on the application of thevascularized three dimensional cardiac construct and the type of cardiactissue material that is desired, the above types of cells may be usedindependently or in combination. In one embodiment, the vascularizedthree dimensional cardiac construct may be composed of autologousprimary tissue isolates from the heart of a patient. Alternatively,cells such as non-immunogenic universal donor cell lines or stem cellsmay be used.

In another embodiment of the present invention, a vascularized threedimensional engineered ligament construct is generated. The cell typesthat may be used to generate a three dimensional ligament construct mayinclude, without limitation, tenocytes, ligamentum cells, fibroblasts,chondrocytes, and endothelial cells. These cells may be usedindependently or in combination and may be primary cells or derived fromcell lines.

Vascularized tissue engineered constructs generated using the methods ofthe present invention have various in vitro and in vivo biomedicalapplications

In one embodiment, a tissue engineered construct of the presentinvention is employed in an in vitro method of screening a test agent.In vitro screening may include, without limitation, toxicologicaltesting of an agent, drug discovery, and biological and chemical warfaredetection. Suitable drugs or agents to be tested using the tissueconstructs of the present invention include cytotoxic agents,pharmaceutical agents, growth factors, etc. In accordance with thisaspect of the invention, a test agent is contacted with a vascularizedtissue construct of the present invention and one or more biologicalendpoints is assayed. Suitable biological endpoints to be assayedinclude, without limitation, carcinogenicity, cell death, cellproliferation, gene expression, protein expression, cellular metabolism,and any combination thereof. Because cellular spatial organization andneovascularization can be induced and controlled in the in vitro culturesystem of the present invention, the resulting vascularized engineeredtissue construct better replicates the in vivo tissue architecture andcellular microenvironment, providing an improved in vitro test model.Use of vascularized engineered tissue constructs of the presentinvention provides an attractive alternative to the use of animal modelsfor testing and screening agents.

In another embodiment, the vascularized tissue engineered constructsgenerated using the methods of the present invention are suitable forimplantation and transplantation into a recipient subject in need oftissue repair or tissue replacement. The method involves selecting asubject that is in need of tissue repair or tissue replacement andimplanting a suitable vascularized engineered tissue construct of thepresent invention into the selected subject. In accordance with thisaspect of the present invention, a suitable subject is any animal,preferably a mammal, more preferably, a human subject.

The vascularized engineered tissue constructs of the present inventionmay be used to replace or augment existing tissue, to introduce new oraltered tissue, to modify artificial prostheses, or to join togetherbiological tissues or structures. For implantation or transplantation invivo, either the cells obtained from the in vitro culture system or,more preferably, the entire vascularized three dimensional engineeredtissue construct is implanted, depending on the type of tissue involved.In accordance with this aspect of the invention, it is desirable to useallogeneic cells in the in vitro culture system. Allogeneic cells ortissue originate from or is derived from a donor of the same species asthe recipient.

The following examples illustrate various methods for compositions inthe treatment method of the invention. The examples are intended toillustrate, but in no way limit, the scope of the invention.

EXAMPLES Materials and Methods for Examples 1-7

Experimental Set-Up.

The experimental set-up used for all USWF exposures is depicted in FIG.1A. A plastic exposure tank (36×20×18 cm) was filled with degassed,deionized water at room temperature. The acoustic source consisted of a1 MHz unfocused transducer, fabricated from a 2.5 cm diameterpiezoceramic disk. The transducer was mounted on the bottom of the watertank. The signal driving the transducer was generated by a waveformgenerator (Model 33120A, Hewlett Packard, Palo Alto, Calif., USA), RFpower amplifier (Model 2100L, ENI, Rochester, N.Y., USA), and anattenuator (Model 837, Kay Elemetrics Corp., Lincoln Park, N.J., USA).Samples were contained within the wells of a modified siliconeelastomer-bottomed cell culture plate (BioFlex® culture plates, FlexCellInternational Corporation, Hillsborough, N.C., USA). These sampleholders were mounted to a three-axis positioner (Series B4000 Unislide,Velmex Inc., East Bloomfield, N.Y., USA) to allow precise control overtheir location within the sound field. The air interface above thesamples was used as the acoustic reflector to generate an USWF withinthe sample volume.

Sample Holder Preparation.

The BioFlex® culture plates used as sample holders for theseinvestigations are depicted in FIG. 1B. They were modified from themanufacturer's form by reducing the diameter of 3 wells per plate from 4cm to 1 cm using Sylgard® 184 Silicone Elastomer (Dow CorningCorporation, Midland, Mich., USA). Through this modification, thediameter of the sample was comparable in size to the width of theultrasound beam. The two-part silicone elastomer was mixed in a 10:1ratio as recommended by the manufacturer's instructions. The solutionwas degassed at room temperature using a vacuum chamber (Model 5830,National Appliance Company, Portland, Oreg., USA) and was subsequentlypoured around 1 cm diameter Teflon® mandrels (Dupont, Wilmington, Del.,USA) that were placed at the center of the 3 wells of interest.Following curing of the silicone elastomer at 20° C. for 48 hr, themandrels were carefully removed to leave a 1 cm diameter sample spacewithin 3 wells of each BioFlex® culture plate (FIG. 1B).

Attenuation Measurements.

The acoustic attenuations of the silicone elastomer well bottom of theBioFlex® plates, the Sylgard® 184 Silicone Elastomer, and standardtissue culture polystyrene (Corning/Costar, Cambridge, Mass., USA) weremeasured using an insertion loss technique. Using the water tank set-up,each material was inserted into the acoustic path between the unfocused2.5 cm diameter, 1 MHz transducer and a hydrophone (either a bilaminarPVDF membrane hydrophone (Marconi Research Center, Chelmsford, England)or a ceramic-based needle hydrophone (Model HNC-0400, Onda Corporation,Sunnyvale, Calif., USA)). Peak positive and peak negative pressureamplitudes were measured using the hydrophone and a digital oscilloscope(Model 9310AM, LeCroy, Chestnut Ridge, N.Y., USA) in the presence andabsence of each material for various source amplitudes. The thickness ofeach material was measured using calipers. The acoustic attenuationcoefficient (in dB/MHz/cm) was calculated for each material.

Absorption/Heating Measurements.

The acoustic absorption coefficient of Sylgard® 184 Silicone Elastomerwas measured using a thermocouple technique. Briefly, a 50 μmcopper-constantan thermocouple was embedded in a sample of Sylgard® 184Silicone Elastomer. Using the water tank set-up, the active element ofthe embedded thermocouple was positioned at the focus of a 1 MHztransducer fabricated from a 3.8 cm diameter plane, piezoceramic diskcemented to the back of a plano-concave lens. A laboratory thermometer(Model BAT-4, Bailey Instruments Co. Inc., Saddle Brook, N.J., USA) anddigital oscilloscope were used to monitor the thermocouple output forvarious pulsing parameters and exposure amplitudes. For each exposurecondition, the initial rate of temperature rise in the sample and thespatial peak temporal average intensities (I_(spta)) were measured andused to calculate the absorption coefficient. The calculated absorptioncoefficients from each exposure condition were averaged to determine theacoustic absorption coefficient (in dB/cm) of Sylgard® 184 SiliconeElastomer at 1 MHz.

Temperature changes in the collagen/cell samples were also monitoredduring USWF exposure using a 50 μm copper-constantan thermocouple.Thermocouple output was monitored using a digital laboratory thermometer(Model BAT-12, Physitemp Instruments Inc., Clifton, N.J., USA),sensitive to changes of 0.1° C., over the duration of USWF exposures.

Acoustic Field Measurements.

Using the water tank set-up, axial spatial distributions of pressurefrom the 1 MHz, 2.5 cm diameter unfocused transducer were measured underUSWF exposure conditions in both the presence and absence of the sampleholder. The Onda ceramic-based needle hydrophone, connected to athree-axis positioner, and a digital oscilloscope were used to measurethe acoustic pressure. The sample holder was placed in the far-fieldwith the well bottoms situated at an axial distance of 12.2 cm from thetransducer. Axial spatial distributions of pressure were measuredthrough a 0.5 cm distance below the air interface in 0.1 mm intervals. Asinusoidal pulse of 50 μs duration was employed and peak positivepressures were measured for each position. The 0.5 cm distanceapproximates the height of the collagen samples used in theseinvestigations. At the axial distance of 12.2 cm from the transducer,the −6 dB transaxial beamwidth in the free field was measured to be 1.2cm.

Acoustic Field Calibrations.

Prior to each experiment, the acoustic field was calibrated using eitherthe Marconi PVDF membrane hydrophone or the Onda ceramic-based needlehydrophone under traveling wave conditions. Hydrophones were calibratedregularly using the steel sphere radiometer technique (Dunn et al., “APrimary Method for the Determination of Ultrasonic Intensity WithElastic Sphere Radiometer,” Acustica 388:58-61 (1977), which is herebyincorporated by reference in its entirety). Acoustic pressure wasmeasured in the far-field at an axial distance of 12.2 cm from thetransducer (where samples were located during USWF exposure).Coordinates from the exposure site to a fixed pointer were determinedusing the three-axis positioner and were used to position the center ofthe lower, left-hand well of the sample holder at the exposure site(bottom of the well was 12.2 cm from the transducer). Some water wasremoved from the tank such that the sample holder was located at theexposure site without full submersion.

Cell Culture.

Fibronectin-null mouse embryonic myofibroblasts (obtained from Dr. JaneSottile, University of Rochester) were used for all experiments. Thesecells do not produce fibronectin and have been adapted to grow underserum-free conditions (Sottile et al., “Fibronectin Matrix AssemblyEnhances Adhesion-dependent Cell Growth,” J Cell Sci 111:2933-43 (1998),which is hereby incorporated by reference in its entirety). Cells wereroutinely cultured in a 1:1 mixture of AimV (Invitrogen, Carlsbad,Calif., USA) and Cellgro (Mediatech, Herndon, Va., USA) on tissueculture dishes pre-coated with collagen type-I. These media do notrequire serum supplementation. Thus, no source of fibronectin is presentduring routine culture. On the day of USWF exposure, fibronectin-nullcells were harvested from monolayer culture by treatment with 0.08%trypsin (Invitrogen) and 0.5 mM EDTA in PBS. Trypsin activity wasneutralized with 2 mg/ml soybean trypsin inhibitor (STI; Sigma, St.Louis, Mo.). Cells were washed one time with 1 mg/ml STI in PBS and werethen resuspended in a 1:1 mixture of AimV/Cellgro.

Collagen Solution Preparation.

A neutralized type-I collagen solution was prepared on ice by mixingcollagen type-I, isolated from rat tail tendons, with 2× concentratedDulbecco's modified Eagle's medium (DMEM; Invitrogen) and 1× DMEMcontaining HEPES so that the final mixture consisted of 0.8 mg/mlcollagen and 1×DMEM (Hocking et al., “Stimulation of Integrin-mediatedCell Contractility by Fibronectin Polymerization,” J Biol Chem275:10673-82 (2000), which is hereby incorporated by reference in itsentirety). Both the 1× and 2×DMEM media were degassed in a vacuumchamber for 30 min under sterile conditions prior to incorporation intothe collagen mixture.

USWF Exposures.

Fibronectin-null cells were added to aliquots of neutralized type-Icollagen solutions on ice at various final concentrations immediatelyprior to USWF exposure. Aliquots (400 μl) of the collagen/cell solutionwere then loaded into two of the 1 cm diameter Sylgard® 184 SiliconeElastomer molded wells of the BioFlex® plate. For “no-cell” samples, anequal volume of AimV/Cellgro was added in place of fibronectin-nullcells and aliquots were loaded into a third well. The collagen/cellsolution in the left-hand well of each plate was exposed to a 1 MHz,continuous wave USWF for 15 min at room temperature. The two othersamples in the plate (right-hand side) served as sham control wells thatwere treated exactly as the exposed sample but were not exposed to theUSWF. The 15 min exposure duration was sufficient to promote collagenpolymerization at room temperature. Following USWF exposure, collagengels were incubated for 1 hr at 37° C. and 8% CO₂ to allow for completecollagen polymerization. An equal volume (400 μl) of DMEM was then addedto wells containing collagen gels. In some experiments, collagen/celland collagen/no-cell solutions were incubated for 1 hr at 37° C. and 8%CO₂ in the sample holders to allow collagen polymerization before USWFexposure.

Cell Viability Assay.

Thiazolyl blue tetrazolium bromide (MTT) was used to assess cellviability (Mosmann T., “Rapid Colorimetric Assay for Cellular Growth andSurvival: Application to Proliferation and Cytotoxicity Assays,” JImmunol Methods 65:55-63 (1983), which is hereby incorporated byreference in its entirety). At various time points after USWF exposure,collagen gels were incubated with 5.3 mM MTT (USB Corporation,Cleveland, Ohio, USA) for 4 hr at 37° C. and 8% CO₂. Gels were thendigested with 0.77 mg/ml collagenase (from Clostridium histolyticum,type-I, Sigma) and formazan crystals were dissolved using acidifiedisopropanol (0.04 N HCl). Absorbance measurements at 570 nm and 700 nm(background) were determined using a spectrophotometer. MTT absorbancewas calculated by subtracting background absorbance values andnon-specific reduction of MTT in no-cell gels from the 570 nm readings.There was a linear relationship between cell number and MTT absorbance.This assay is sensitive to differences of 5000 cells and greater.

Collagen Gel Contraction Assays.

The extent of collagen gel contraction was determined using twoestablished methods. For volumetric gel contraction assays, gels werescored around their edges to form free-floating gels in the wells. Afteran additional 20 hr of incubation at 37° C. and 8% CO₂, the gels wereremoved from the wells and weighed (Model B303, Mettler Toledo,Columbus, Ohio, USA). Volumetric collagen gel contraction was calculatedas a decrease in gel weight as compared to the control, no-cell gelweight (Hocking et al., “Stimulation of Integrin-mediated CellContractility by Fibronectin Polymerization,” J Biol Chem 275:10673-82(2000), which is hereby incorporated by reference in its entirety). Forradial gel contraction assays, gel diameters were measured using a 10×inspection microscope equipped with a calibrated eyepiece micrometer.Two measurements were recorded for each gel and averaged to calculategel diameter. Investigators measuring diameters were blinded to exposureconditions. Radial collagen gel contraction was calculated as a decreasein gel diameter as compared to the original gel diameter of 1 cm(Tingstrom et al., “Regulation of Fibroblast-mediated Collagen GelContraction by Platelet-derived Growth Factor, Interleukin-1 Alpha andTransforming Growth Factor-betal,” J Cell Sci 102:315-22 (1992), whichis hereby incorporated by reference in its entirety).

Soluble Fibronectin Binding.

Fibronectin-null cells in suspension (2×10⁷ cell/ml) were incubated with100 μg/ml of Alexa Fluor® 488-labeled human, plasma-derived fibronectin(FN-488; labeled according to manufacturer's instructions) in thepresence of 1 mM MnCl₂ for 30 min at room temperature (Akiyama et al.,“The Interaction of Plasma Fibronectin With Fibroblastic Cells inSuspension,” J Biol Chem 260:4492-500 (1985) and Mastrangelo et al.,“Amino Acid Motifs for Isolated Beta Cytoplasmic Domains to Regulate ‘inTrans’ Beta-1 Integrin Conformation and Function in Cell Attachment,” JCell Sci 112:217-29 (1999), which are hereby incorporated by referencein their entirety). Cells were washed twice with AimV/Cellgro to removeunbound fibronectin and were then added to neutralized type-I collagensolutions and exposed to an USWF as described above. In otherexperiments, 10 μg/ml of FN-488 was added to neutralized type-I collagensolutions in the absence of cells and exposed to an USWF as describedabove.

Microscopy.

One hour after USWF exposure, cell-embedded collagen gels were examinedusing an Olympus IX70 inverted microscope (Center Valley, Pa., USA) witha 4× phase-contrast objective and were photographed using a digitalcamera (Spot RT Slider, Model 2.3.1, Diagnostic Instruments Inc.,Sterling Heights, Mich., USA). FN-488 was visualized usingepifluorescence microscopy. Gels were flipped on their side to visualizecell bands through the height of the cylindrical sample. For volumetriccollagen gel contraction experiments, gels were imaged after obtainingweight data. Image-Pro Plus software (Media Cybernetics, Bethesda, Md.,USA) was used to measure the linear distance between fibronectin-nullcell bands within collagen gels. Pixel distance was converted to micronvalues using a micrometer calibration. A total of 10 distances weremeasured on each of 20 different images collected from 3 differentexperiments.

To visualize type-I collagen fibers, cell-embedded collagen gels wereexamined using second-harmonic generation microscopy (Freund et al.,“Second-harmonic Microscopy of Biological Tissue,” Opt Lett 11:94-6(1986); Roth et al., “Second Harmonic Generation in Collagen,” J ChemPhys 70:1637-43 (1979); and Williams et al., “InterpretingSecond-Harmonic Generation Images of Collagen I Fibrils,” Biophys J88:1377-86 (2005), which are hereby incorporated by reference in theirentirety). One hour after USWF exposure, gels were fixed in 4%paraformaldehyde for 1 hr at room temperature. Second-harmonicgeneration microscopy was performed using an Olympus Fluoview 1000AOM-MPM microscope equipped with a 25×, 1.05 NA water immersion lens(Olympus). Samples were illuminated with 780 nm light generated by a MaiTai HP Deep See Ti:Sa laser (Spectra-Physics, Mountain View, Calif.,USA) and the emitted light was detected with a photomultiplier tubeusing a bandpass filter with a 390 nm center wavelength (FilterFF01-390/40-25, Semrock, Inc., Rochester, N.Y., USA). Fibronectin-nullcells were simultaneously visualized using a second bandpass filter witha 519 nm center wavelength (Filter BA 495-540 HQ from MPFC1, Olympus) byexploiting the intrinsic auto-fluorescence of cells (Monici M., “Celland Tissue Autofluorescence Research and Diagnostic Applications,”Biotechnol Annu Rev 11:227-56 (2005), which is hereby incorporated byreference in its entirety). Cell-embedded collagen gels werephotographed using a CMOS digital camera (Moticam 1000, Motic, China).

Statistical Analyses.

Data are presented as the mean□±SEM. Statistical comparisons betweenUSWF-exposed and sham experimental conditions were performed usingeither the Student's t test for paired samples or one-way analysis ofvariance in GraphPad Prism software (La Jolla, Calif., USA). Differenceswere considered significant for p values <0.05.

Example 1—Experimental Set-Up for Ultrasound Exposures

To investigate the effects of ultrasonic mechanical forces on thepromotion of angiogenesis, an ultrasound (US) exposure system wasdeveloped in which cells and 3D tissue constructs are subjected to anultrasound traveling wave field (UTWF) or an ultrasound standing wavefield (USWF) (FIG. 1A). Samples are contained within the wells of amodified silicone elastomer-bottomed cell culture plate (FIG. 1B). UTWFare created by minimizing reflections at the sample/air interface with arubber absorber. USWF are produced by removing the rubber absorber suchthat the air interface promotes the interference of incident andreflected waves (Blackstock DT, FUNDAMENTALS OF PHYSICAL ACOUSTICS,(Wiley & Sons, 2000), which is hereby incorporated by reference in itsentirety). No significant differences in UTWF or USWF are found in thepresence of the sample holder compared to without the holder, indicatingthat the experimental set-up will not interfere with the sound field.

The ultrasound attenuation of standard polystyrene multi-well tissueculture plates was measured to be 4.5±0.7 □dB/MHz/cm (n=3). Due to thesignificant attenuation of the sound field by these polystyrene plates,the use of silicone elastomer-bottomed plates was investigated as analternative sample holder for these studies. The acoustic attenuation ofthe silicone elastomer well bottom (thickness=1 mm) of the BioFlex®plates was measured to be only 0.6±0.4 dB/MHz/cm (n=3) indicating thatthere is negligible attenuation (0.06 dB at 1 MHz) of the sound fielddue to the presence of the BioFlex® sample holders.

The ultrasound attenuation of the Sylgard® 184 Silicone Elastomermolding material was measured to be 2.4±0.04 dB/MHz/cm (n=3). Soundabsorption at 1 MHz (1.4±0.03 dB/cm; n=3) was found to contribute to˜60% of this attenuation. Thermocouple measurements monitoring sampletemperature during USWF exposure indicated that collagen/cell sampletemperatures never exceeded that of room temperature. Therefore, theBioFlex® plates modified with the Sylgard® 184 Silicone Elastomer moldswere chosen as sample holders for these investigations because they didnot significantly interfere with the sound field.

Example 2—Characterization of Ultrasound Fields

USWF and traveling wave fields were measured in a water tank and withinthe sample space using the set-up described above. Resulting spatialdistributions of pressure are illustrated below in FIGS. 2A-2D. Anunfocused, 1 MHz, 1 inch diameter, piezoelectric transducer was used toproduce the US field and a needle hydrophone was used to measure theacoustic pressure.

For traveling wave fields, transaxial spatial distributions in pressurewere measured in the far field, 12.2 cm from the transducer. In theabsence of the sample holder, −3 dB and −6 dB beam widths were 0.8 cmand 1.2 cm, respectively (FIG. 2A). The beam pattern within the samplespace was not significantly altered from the free field patternindicating that the sample holder does not interfere with soundpropagation (FIG. 2B). The −3 dB beam width remained 0.8 cm, but the −6dB beam width could not be calculated, indicating that samples will beexposed to a relatively uniform pressure distribution within the −6 dBbeam width.

For standing wave fields, axial spatial distributions in pressure weremeasured through a 1 cm distance, starting at the air interface. In boththe absence

(FIG. 2C) and presence (FIG. 2D) of the sample holder, beam patternsexhibited characteristic pressure maxima and minima indicating that thesample holder did not interfere with the development of the USWF. Adistance of 0.8 mm separated pressure minima in both beam patterns, afinding that is consistent with the expected half-wavelength spacingbetween the pressure nodes in an USWF at 1 MHz.

Example 3—USWF Control of the Spatial Arrangement of Cells Within a 3DTissue Construct

The organization of endothelial cells into multicellular assembliesaffects angiogenic endothelial cell behaviors (Korff et al.,“Integration of Endothelial Cells in Multicellular Spheroids PreventsApoptosis and Induces Differentiation,” The Journal of Cell Biology143(5):1341-1352 (1998) and Ino et al., “Application of MagneticForce-Based Cell Patterning for Controlling Cell-Cell Interactions inAngiogenesis,” Biotechnology and Bioengineering 102(3):882-890 (2009),which are hereby incorporated by reference in their entirety). Exposureof cell suspensions to USWF can result in cellular aggregation at areasof minimum acoustic pressure (the pressure nodes) (Dyson et al., “TheProduction of Blood Cell Stasis and Endothelial Damage in the BloodVessels of Chick Embryos Treated with Ultrasound in a Stationary WaveField,” Ultrasound in Medicine and Biology 1:133-148 (1974), which ishereby incorporated by reference in its entirety). An USWF acousticradiation force is largely responsible for this cell movement (Coakleyet al., “Cell Manipulation in Ultrasonic Standing Wave Fields,” Journalof Chemical Technology and Biotechnology 44:43-62 (1989), which ishereby incorporated by reference in its entirety). To determine if USWFradiation forces could manipulate cell organization in 3D collagen gels,fibronectin-null myofibroblasts (FN−/− MF), suspended in unpolymerizedtype-I collagen solutions, were either exposed to, or not exposed to(sham samples), a 1 MHz, continuous wave (CW) USWF using theexperimental set-up shown in FIG. 1A. Collagen solutions were allowed topolymerize during the 15 min exposure to maintain the US-induced celldistribution after removal of the pressure field (Saito et al.,“Composite Materials with Ultrasonically Induced Layer or LatticeStructure,” Jpn Journal of Applied Physics 38:3028-3031 (1999); Saito etal., “Fabrication of a Polymer Composite with Periodic Structure by theUse of Ultrasonic Waves,” Journal of Applied Physics 83(7):3490-3494(1998); Gherardini et al., “A New Immobilisation Method to ArrangeParticles in a Gel Matrix by Ultrasound Standing Waves,” Ultrasound inMedicine and Biology 31(2):261-272 (2005); and Gherardini et al., “AStudy of the Spatial Organization of Microbial Cells in a Gel MatrixSubjected to Treatment with Ultrasound Standing Waves,” Bioseparation10:153-162 (2002), which are hereby incorporated by reference in theirentirety). Cell distribution was then analyzed using phase-contrastmicroscopy (FIGS. 3A-3C). FN−/− MF do not produce fibronectin and aregrown in serum-free conditions (Sottile et al., “Fibronectin MatrixAssembly Enhances Adhesion-Dependent Cell Growth,” Journal of CellScience 111:2933-2943 (1998), which is hereby incorporated by referencein its entirety). As such, these cells were chosen for the initialstudies to differentiate between the effects of USWF on the localizationof cells and the extra-cellular matrix protein fibronectin. Gelspolymerized in the presence of the USWF showed a distinct bandedpattern, characterized by the localization of cells to the pressurenodes of the USWF (FIGS. 3B-3C), while sham samples exhibited ahomogeneous cell distribution (FIG. 3A). The mean of the measureddistance between cell bands was 657±15 μm. This is consistent with theexpected half-wavelength spacing of 750 μm between pressure nodal planesin an USWF generated with a 1 MHz source. Increasing the initialconcentration of cells in the collagen solutions leads to the formationof denser cell bands at the nodal planes (compare FIGS. 3B and 3C).These data indicate that an USWF can spatially organize cells within 3Dcollagen gels, and that the extent of the banded pattern of cells isdependent on cell concentration.

In supplementary experiments, the MTT assay (Mosmann T., “RapidColorimetric Assay for Cellular Growth and Survival: Application toProliferation and Cytotoxicity Assays,” Journal of Immunological Methods65(1-2):55-63 (1983), which is hereby incorporated by reference in itsentirety) was used to show that USWF exposure did not adversely affectcell viability (FIG. 4). These findings indicate that USWF cannoninvasively alter the spatial organization of cells within a 3Dcollagen gel without affecting cell viability.

To estimate the magnitude of radiation force exerted on the cells in theapplied USWF, Equation 1 was used to calculate the maximum F_(rad). Theacoustic exposure parameters and the physical properties of the cellsand the suspending collagen medium used for the calculation are listedin Table 1. Results of this calculation indicated that the cells weresubjected to a maximum radiation force of approximately 2.2 pN.

TABLE 1 Parameters used for calculation of primary acoustic radiationforce (F_(rad)) using Equation 1 Parameters (units) Numerical ValueSource/Reference P_(o) (MPa) 0.2 Hydrophone measurement V (μm³) 904.8(cells) Assuming spherical particles 4.2 × 10⁻⁶ (FN) r (μm) 6 (cells)cell radius measured for 0.01 (FN) rounded fibronectin-null cells insuspension; FN radius (Vuillard et al. 1990) β_(o) (1/Pa) 4.44 × 10⁻¹⁰Calculation from β = 1/c²ρ β_(p) (1/Pa) 4.07 × 10⁻¹⁰ (cells) 3.12 ×10⁻¹⁰ (FN) c_(o) (m/s) 1500 Assuming collagen media ρ_(o) (kg/m³) 1000has properties of water λ (μm) 1500 at room temperature f (MHz) 1 Chosenfrequency for these studies φ 0.132 (cells) Using Equation 2 0.581 (FN)c_(p) (m/s) 1529 (cells) cell value (Taggart et al. 2007); 1540 (FN) FNvalue assumed to be sound speed in human soft tissue (Bamber 1998) ρ_(p)(kg/m³) 1050 (cells) cell value assumed to be 1350 (FN) dominated bycytoplasm taken as low concentration saline (Baddour et al. 2005); FNvalue (Fischer et al. 2004) z (μm) 0-750 axial distance between 2pressure nodal planes (λ/2) F_(rad) max (pN) ±2.2 (cells) Calculatedusing Equation 1 ±4.5 × 10⁻⁸ (FN) at z = λ/8 and 3λ/8 References citedin Table 1 which are hereby incorporated by reference in their entirety:Baddour et al., “High-Frequency Ultrasound Scattering from Microspheresand Single Cells,” J Acoust Soc Am 117: 934-43 (2005); Bamber JC,Ultrasonic Properties of Tissues, in ULTRASOUND IN MEDICINE 57-88(1998); Fischer et al., “Average Protein Density is aMolecular-Weight-Dependent Function:” Protein Sci 13: 2825-8 (2004);Taggart et al., “Ultrasonic Characterization of Whole Cells and IsolatedNuclei,” Ultrasound Med Biol 33: 389-401 (2007)

Example 4—USWF Radiation Forces Control Soluble FN Organization Within3D Constructs

Successful tissue engineering depends upon the stimulation of key cellfunctions, including cell proliferation, migration and differentiation.These processes are influenced by a variety of soluble and insolublefactors, including growth factors, cytokines, and extracellular matrixproteins (Langer and Vacanti, “Tissue Engineering,” Science 260:920-6(1993), which is hereby incorporated by reference in its entirety).Concentrating stimulatory proteins to the cell banded areas of collagengels may stimulate cell function. The extracellular matrix protein,fibronectin, stimulates cell growth, migration, and contractility(Hocking et al., “Stimulation of Integrin-Mediated Cell Contractility byFibronectin Polymerization,” J Biol Chem 275:10673-82 (2000); Hockingand Chang, “Fibronectin Polymerization Regulates Small Airway EpithelialCell Migration,” Am J Physiol Lung Cell Mol Physiol 285:L169-L79 (2003);Sottile et al., “Fibronectin Matrix Assembly Enhances Adhesion-DependentCell Growth,” J Cell Sci 111:2933-43 (1998), which are herebyincorporated by reference in their entirety).

USWF radiation force theory predicts that small protein molecules suchas fibronectin will not localize to the pressure nodes of an USWF(Coakley et al., “Cell Manipulation in Ultrasonic Standing Wave Fields,”Journal of Chemical Technology and Biotechnology 44:43-62 (1989) andWhitworth et al., “Particle Column Formation in a Stationary UltrasonicField,” Journal of the Acoustical Society of America 91(1):79-85 (1992),which are hereby incorporated by reference in their entirety). Using theaforementioned experimental set-up and USWF exposure conditions, thisidea was confirmed by including Alexa488-labeled, human, plasma-derivedfibronectin (FN-488) in unpolymerized collagen solutions and allowingpolymerization to occur during USWF exposure. Fluorescent microscopyimages show a homogeneous distribution of soluble fibronectin remainingafter USWF exposure (FIG. 5A bottom left panel). To facilitatelocalization of fibronectin to cell-aggregated areas of collagen gels,soluble FN-488 molecules were bound to FN−/−MF prior to USWF exposure.Fluorescent microscopic analysis shows a co-localization of fibronectinmolecules to USWF-induced cell bands within the collagen gels (FIG. 5Bbottom right panel) indicating that USWF radiation forces cannoninvasively control the organization of cell-bound fibronectin within3D tissue constructs. As such, the mechanical forces associated withultrasound can influence extracellular matrix (ECM) organization andthus have the potential to affect endothelial cell functions essentialto the angiogenic process.

Example 5—USWF-Induced Cell Organization Enhances Cell-Mediated CollagenGel Contraction

The biomechanical properties of normal human tissue and engineeredtissue constructs are in part dictated by the organization of theextracellular matrix comprising that tissue or tissue construct (Vogelet al., “Local Force and Geometry Sensing Regulate Cell Functions,” NatRev Mol Cell Biol 7:265-75 (2006), which is hereby incorporated byreference in its entirety). In turn, extracellular matrix organizationis partly influenced by cell-derived forces. These forces are exerted onmatrix components through intracellular tension generation due tocytoskeletal contractility (Hinz et al., “Mechanisms of Force Generationand Transmission by Myofibroblasts,” Curr Opin Biotechnol 14:538-46(2003); Hocking et al., “Stimulation of Integrin-mediated CellContractility by Fibronectin Polymerization,” J Biol Chem 275:10673-82(2000); and Lee et al., “Extracellular Matrix and PulmonaryHypertension-control of Vascular Smooth Muscle Cell Contractility,” Am JPhysiol Heart Circ Physiol 274:H76-H82 (1998), which are herebyincorporated by reference in their entirety). Cell-mediated collagen gelcontraction is a common measure of extracellular matrix remodeling bycells (Korff et al., “Tensional Forces in Fibrillar ExtracellularMatrices Control Directional Capillary Sprouting,” J Cell Sci112:3249-58 (1999); Sieminski et al., “The Relative Magnitudes ofEndothelial Force Generation and Matrix Stiffness Modulate CapillaryMorphogenesis In Vitro,” Exp Cell Res 297:574-84 (2004); and Vernon etal., “Contraction of Fibrillar Type I Collagen by Endothelial Cells: AStudy In Vitro,” J Cell Biochem 60:185-97 (1996), which are herebyincorporated by reference in their entirety). Changes in the extent ofcollagen gel contraction indicate alterations in ECM remodeling that cansubsequently influence cell behavior (Sieminski et al., “The RelativeMagnitudes of Endothelial Force Generation and Matrix Stiffness ModulateCapillary Morphogenesis In Vitro,” Experimental Cell Research297:574-584 (2004); Vernon et al., “Reorganization of Basement Membraneby Cellular Traction Promotes the Formation of Cellular Networks InVitro,” Laboratory Investigation 66:536-547 (1992); Korff et al.,“Tensional Forces in Fibrillar Extracellular Matrices ControlDirectional Capillary Sprouting,” Journal of Cell Science 112:3249-3258(1999); Vernon et al., “Contraction of Fibrillar Type I Collagen byEndothelial Cells: A Study In Vitro,” Journal of Cellular Biochemistry60:185-197 (1996), which are hereby incorporated by reference in theirentirety). To assess if a change in the spatial distribution of cellsaffects cell-mediated matrix remodeling, a collagen gel contractionassay was used to compare the extent of collagen gel contraction betweenUSWF cell-organized and sham gels. A 2-fold increase in the contractionof collagen gels with USWF-induced cell organization was found ascompared to sham gels with a random cell distribution (FIGS. 6A and 6B).Additional experiments were designed to demonstrate that this increasein collagen gel contraction was a result of the altered celldistribution and not an effect of the USWF exposure parameters on FN−/−MF tension generation. According to theory, increasing the viscosity ofthe suspending medium will inhibit USWF-induced movement of cells to thepressure nodes (Coakley et al., “Cell Manipulation in UltrasonicStanding Wave Fields,” Journal of Chemical Technology and Biotechnology44:43-62 (1989), which is hereby incorporated by reference in itsentirety). Based on this prediction, collagen gels with a homogeneousdistribution of FN−/− MF were polymerized prior to USWF exposure. Nodifferences in collagen gel contraction between USWF-exposed and shamsamples were found and microscopic analysis confirmed the lack oflocalization of cells to the nodal planes (FIGS. 6C and 6D). These dataindicate that cell-mediated ECM remodeling is altered in USWFcell-organized collagen gels.

Example 6—The Formation of Cell Bands Depends on USWF Pressure Amplitude

The magnitude of the acoustic radiation force (F_(rad)) in an USWF has asecond order dependence on pressure amplitude (P_(o)) and, as such,changing P_(o) will affect the movement of cells to the pressure nodes(Eq. 1). Theoretical analysis of the forces acting on particles in anUSWF predicts a threshold pressure for banding below which particleswill not accumulate on the nodal planes (Coakley et al., “CellManipulation in Ultrasonic Standing Wave Fields,” J Chem Tech Biotechnol44:43-62 (1989), which is hereby incorporated by reference in itsentirety). To determine the threshold pressure amplitude necessary toachieve cell banding within collagen gels, fibronectin-null cellssuspended in type-I collagen solutions were exposed during thepolymerization process to an USWF of various peak pressure amplitudes.Cell distribution was then analyzed using phase-contrast microscopy. Asshown in FIG. 7, homogeneous cell distributions were observed withinsham-exposed gels, as well as gels fabricated using an USWF with peakpressure amplitudes of 0.02 and 0.05 MPa. Exposing samples to an USWFwith peak pressure amplitude of 0.1 MPa resulted in the formation ofcell bands, indicating that the threshold pressure for USWF-induced cellbanding in this system is ˜0.1 MPa (F_(rad)=0.55 pN). When an USWF withpressure amplitude of 0.2 MPa is used to fabricate the collagen gels,cell bands appeared more dense. With a pressure amplitude of 0.3 MPa,the resulting cell bands were thicker and more localized to the centerof the gel, likely due to the influence of secondary lateral acousticradiation forces acting within the pressure nodal planes (Spengler etal., “Microstreaming Effects on Particle Concentration in an UltrasonicStanding Wave,” AIChE J 49:2773-82 (2003), which is hereby incorporatedby reference in its entirety). These findings indicate that differentUSWF pressure amplitudes lead to variations in the patterns of bandedcells within collagen gels.

Example 7—USWF Pressure Amplitude Has a Biphasic Effect on Cell-MediatedCollagen Gel Contraction

The data indicate that USWF-induced cell organization enhancescell-mediated collagen gel contraction, and that the extent of cellbanding is affected by the USWF pressure amplitude. To determine ifdifferent cell banded patterns affect the extent of collagen gelcontraction, radial collagen gel contraction assays were used to comparelevels of collagen gel contraction among gels fabricated at the sixdifferent USWF pressure amplitudes shown in FIG. 7. No differences incollagen gel contraction were observed among sham-exposed samples andsamples exposed to either 0.02 or 0.05 MPa (FIG. 8A) where cells remainin a homogeneous distribution (FIG. 7). In contrast, a significant1.5-fold increase in gel contraction occurred at 0.1 MPa (FIG. 8A), thepressure threshold for cell banding, where cells first become alignedinto planar bands (FIG. 7). These results, obtained using gel diametermeasurements, are similar to those reported in FIG. 6 using volumetriccontraction assays, and therefore, provide additional evidence thatUSWF-induced cell banding enhances cell-mediated collagen gelcontraction and matrix reorganization.

As the USWF pressure amplitude was increased above 0.1 MPa, collagen gelcontraction levels decreased (FIG. 8A). At 0.3 MPa, a 30% decrease incontraction as compared to sham levels was observed (FIG. 8A). Therewere no significant differences in the number of viable cells betweensham-exposed and 0.3 MPa USWF-exposed samples (FIG. 8B). Therefore, thedecrease in collagen gel contraction was not due to cell death, but wasmore likely due to effects of the cell banded pattern that occurred at0.3 MPa. These findings indicate that the effect of USWF pressureamplitude on cell-mediated collagen gel contraction is biphasic. Thisbiphasic effect on collagen gel contraction can be attributed to thedecrease in cell-extracellular matrix contacts formed as cell bandsbecome more compact above the threshold pressure.

To directly assess collagen matrix organization relative to the cellbanded areas formed using USWF of various pressure amplitudes,second-harmonic generation microscopy imaging of collagen fibers wasperformed. As shown in FIG. 9, short collagen fibrils were randomlyorganized in sham-exposed (0 MPa) cell-embedded collagen gels and incell-embedded collagen gels exposed to either 0.02 or 0.05 MPa wherecells remain in a homogeneous distribution. Exposure to 0.1 MPa resultedin areas of the gels where cells were loosely clustered and collagenfibrils were more elongated. These data clearly show that cells alignedinto planar bands at the pressure threshold for cell banding havereorganized their surrounding collagen matrix, and thus, provide furtherevidence that USWF-induced cell banding enhances cell-mediated collagenmatrix remodeling.

Extensive areas of cell bands were clearly visible in collagen gelsexposed to 0.2 or 0.3 MPa, and short collagen fibers surrounding theseareas were randomly oriented. These results indicate that as the USWFpressure amplitude increases beyond the pressure threshold for cellbanding and cell bands become more dense, cell-mediated collagen matrixreorganization decreases. Therefore, the effect of USWF pressureamplitude on cell-mediated collagen matrix reorganization is biphasicand as such, these data both parallel and support the data showing thatUSWF pressure amplitude has a biphasic effect on collagen gelcontraction. Taken together, these data indicate that radiation forcesassociated with an USWF can indirectly influence the relative locationof extracellular proteins and thus, can be used to controlextracellular-matrix dependent functions essential to tissue formation.

Discussion of Examples 1-7

Examples 1-7 above describe the development and use of ultrasoundstanding wave fields as a novel, non-invasive technology for organizingcells and cell-bound proteins within tissue engineered biomaterials.These studies have demonstrated that acoustic radiation forcesassociated with an USWF can be used to organize both mammalian cells andcell-associated proteins into discrete bands within collagen hydrogels.The density of the USWF-aligned cell bands was dependent on both cellnumber and pressure amplitude. Exposure of cells to USWF parametersutilized in the current study did not decrease cell viability.Furthermore, the USWF-aligned cell bands were stable for at least 20 hr.

Under appropriate conditions, the organization of cells into bands ledto an increase in cell-mediated collagen gel contraction, as measured byboth volumetric and radial changes, demonstrating an increase in cellfunction in response to cell alignment. The increase in collagen gelcontraction in response to USWF exposure did not occur if thecollagen/cell samples were allowed to polymerize prior to USWF exposure,strongly suggesting that the increases in cell contractility andcollagen fibril reorganization were mediated by the organization ofcells into bands and were not an indirect effect of ultrasound exposureon individual cells. The extent of collagen contraction was dependentupon the spatial distribution of cells in the gel. No increase incollagen gel contraction was observed in response to USWF exposure atpressure amplitudes that did not produce cell banding. At the otherextreme, no increase in collagen gel contraction was observed inresponse to USWF at pressure amplitudes that produced densely packedcell bands. However, exposure of samples to an USWF that leads to theclustering of cells into planar bands within the gel resulted in asignificant increase in collagen contraction above sham gels with ahomogenous cell distribution.

Clustering of cells into planar bands in response to an USWF also led tochanges in collagen fibril organization and length. Second-harmonicgeneration microscopic images showed that short collagen fibers wererandomly organized in gels exposed to USWF at pressure amplitudes thatdid not produce cell banding as well as pressure amplitudes thatproduced densely packed cell bands. In contrast, elongated collagenfibers were observed within loosely clustered cell banded areasindicating enhanced cell-mediated collagen matrix remodeling in thesesamples. These results are consistent with the biphasic results of thecollagen gel contraction investigations. Hence, an important downstreameffect of USWF-mediated cell alignment is enhanced extracellular matrixremodeling. Thus, the use of USWF to specifically control cell andextracellular organization is a promising new approach for engineeringcomplex tissues in vitro.

Acoustic radiation forces exerted on extracellular matrix proteins weretoo small to directly organize proteins in the system used in thisstudy. However, the use of an USWF can indirectly effect theorganization of proteins in the extracellular matrix by two avenues.First, as explained in the paragraph above, increased collagen fibrillength and organization was observed in gels exposed to an USWF atpressure amplitudes that increased collagen gel contraction. Thus, adownstream effect of USWF-induced cell banding is the resultant cellularremodeling of the surrounding extracellular matrix. Second, theextracellular matrix protein, fibronectin, could be aligned into bandswithin the collagen gel using an USWF if the fibronectin molecules werefirst bound to the cell surface. Thus, USWF technology can be usedspatially organize cells within engineered tissues and to co-locateactive or inactive cell-bound molecules.

The use of an USWF has numerous advantages as a noninvasive technologyto spatially organize cells and cell-bound molecules in engineeredtissues, and thereby influence cell function. The acoustic radiationforce acts directly on the cells and thus the approach does not requireany prior modification of the cell surface. Various hydrogels thatundergo phase transitions could be adapted to this technique. Changingfrequency of the ultrasound field will affect spacing of cell bands, andmultiple transducers could be used to produce more complex patterns ofcells within engineered biomaterials.

Example 8—Endothelial Cell Sprouting is Observed from USWF-Induced ECBands

Multicellular spheroids of endothelial cells are commonly used to studysprouting angiogenesis in 3D assays (Vailhe et al., “In Vitro Models ofVasculogenesis and Angiogenesis,” Laboratory Investigation 81(4):439-452(2001), which is hereby incorporated by reference in its entirety). Toinvestigate if the organization of EC into a banded pattern withincollagen gels supports endothelial cell sprouting, human umbilical veinendothelial cells (HUVEC) were subjected to an USWF and resultingendothelial cell bands were analyzed for the formation of endothelialcell sprouts. As shown in FIG. 10B, endothelial cells sprouting from acell band can be seen 24 hrs after USWF exposure. Sprouting was absentin sham gels where a rounded cell morphology was found. Thesepreliminary observations suggest that ultrasonic radiation force-inducedendothelial cell organization promotes an angiogenic phenotype inendothelial cell.

Example 9—The Use of Ultrasound to Promote Angiogenesis within 3DCollagen Constructs

Ultrasound-mediated changes in endothelial cell and ECM proteinorganization is expected to induce neovessel formation for thevascularization of 3D tissue constructs. The development of vascularsystems within engineered tissues is essential to the advancement of thetissue engineering field (M.A.T.E.S.I.W.G.

Advancing Tissue Science and Engineering: A Foundation for the Future—AMulti-Agency Strategic Plan. 2007; Griffith et al., “TissueEngineering-Current Challenges and Expanding Opportunities,” Science295:1009-1014 (2002); and Nerem R. M., “Tissue Engineering: The Hope,the Hype, and the Future,” Tissue Engineering 12:1143-1150 (2006), whichare hereby incorporated by reference in their entirety).

New vascular networks can be induced to grow within 3D constructs bystimulating angiogenic endothelial cell behaviors (Nomi et al.,“Principals of Neovascularization for Tissue Engineering,” MolecularAspects of Medicine 23:463-483 (2002); Laschke et al., “Angiogenesis inTissue Engineering: Breathing Life into Constructed Tissue Substitutes,”Tissue Engineering 12(8):2093-2104 (2006); Soker et al., “Systems forTherapeutic Angiogenesis,” World Journal of Urology 18:10-18 (2000); andLokmic et al., “Engineering the Microcirculation,” Tissue Engineering14(1):87-103 (2008), which are hereby incorporated by reference in theirentirety) through the aggregation of endothelial cells intomulticellular structures (Korff et al., “Integration of EndothelialCells in Multicellular Spheroids Prevents Apoptosis and InducesDifferentiation,” The Journal of Cell Biology 143(5):1341-1352 (1998)and Ino et al., “Application of Magnetic Force-Based Cell Patterning forControlling Cell-Cell Interactions in Angiogenesis,” Biotechnology andBioengineering 102(3):882-890 (2009), which are hereby incorporated byreference in their entirety), controlling the organization of thesurrounding ECM (Francis et al., “Endothelial Cell-Matrix Interactionsin Neovascularization,” Tissue Engineering 14(1):19-32 (2008) andSottile J., “Regulation of Angiogenesis by Extracellular Matrix,”Biochimica et Biophysica Acta 1654:13-22 (2004), which are herebyincorporated by reference in their entirety), and exposing endothelialcells to mechanical forces (Iba et al., “Effect of Cyclic Stretch onEndothelial Cells from Different Vascular Beds,” Circulatory Shock35:193-198 (1991); Ando et al., “The Effect of Fluid Shear Stress on theMigration and Proliferation of Cultured Endothelial Cells,”Microvascular Research 33:62-70 (1987); Vouyouka et al., “AmbientPulsatile Pressure Modulates Endothelial Cell Proliferation,” Journal ofMolecular and Cellular Cardiology 30(3):609-615 (1998); which are herebyincorporated by reference in their entirety). The data described supraindicates that USWF radiation forces aggregate cells into multicellulararrangements, organizes fibronectin into a banded pattern, enhancescell-mediated collagen remodeling, and promotes endothelial cellsprouting. These results, together with previous findings thatultrasound can enhance angiogenesis (Barzelai et al., “Low-IntensityUltrasound Induces Angiogenesis in Rat Hind-Limb Ischemia,”Ultrasound inMedicine and Biology 32(1):39-145 (2006); Young et al., “The Effect ofTherapeutic Ultrasound on Angiogenesis,” Ultrasound in Medicine andBiology 16:261-269 (1990); Raz et al., “Cellular Alterations in CulturedEndothelial Cells Exposed to Therapeutic Ultrasound Irradiation,”Endothelium 12:201-213 (2005); Mizrahi et al., “Ultrasound-InducedAngiogenic Response in Endothelial Cells,” Ultrasound in Medicine andBiology 33(11):1818-1829 (2007), which are hereby incorporated byreference in their entirety), suggest that ultrasound can play a role inpromoting an angiogenic response in endothelial cells for the formationof neovascular networks in 3D tissue constructs.

The formation of neovessels in response to USWF-induced endothelial cellorganization was assessed at various times over a 10-day period usingphase-contrast microscopy. Human umbilical vein endothelial cells weresuspended at 1×10⁶ cell/ml in a neutralized type-I collagen solution andexposed to a 1 MHz USWF with a peak pressure amplitude of 0.2 MPa for a15 min duration at room temperature to promote the formation ofmulticellular endothelial cell bands. Cell-embedded collagen gels wereincubated at 37° C. and 5% CO₂ for an additional 10 days. Sham sampleswere treated in the exact same manner as USWF-exposed samples but didnot receive USWF treatment. Representative phase-contrast images of shamand USWF-treated samples were collected on Day 1, 4, 6, 8, and 10 (FIGS.11A-11B; scale bar, 100 μm). Multiple capillary-like sprouts emergedfrom areas of organized endothelial cell bands by Day 1 in USWF-exposedsamples (FIG. 11B, “USWF”). In contrast, endothelial cells were randomlydistributed in sham samples (FIG. 11A, “Sham”). The capillary-likesprouts that formed in USWF-exposed samples increased in length andwidth over the ten-day period (FIG. 11B). In addition, numerous brancheswere observed.

To assess cell morphology within the USWF-induced capillary-likestructures, USWF- and sham-exposed samples were fixed four days afterUSWF exposure, and then processed for immunofluorescence microscopy.Cell nuclei were visualized by staining with DAPI (FIGS. 12A-12B) andhuman umbilical vein endothelial cells were visualized by staining withanti-human CD31 monoclonal antibody followed by AlexaFluor-594conjugated anti-mouse IgG (FIGS. 12A-12B). Two-photon microscopy wasused to collect images along the z-axis in 1 μm slices. Images were thenprojected onto the z-plane using ImageJ software. Endothelial cells insham-exposed samples demonstrated random network formation and condensednuclei, indicative of cell death (FIG. 12A). In contrast, capillarysprouts in USWF-exposed samples were well-organized, multicellularstructures that clearly extended from endothelial cell bands (FIG. 12B,“USWF”; scale bar, 15 microns).

To determine whether lumens were formed within the USWF-inducedcapillary-like structures, USWF- and sham-exposed samples were fixed andprocessed for histological analysis four days after USWF exposure.Four-micrometer thick gel cross-sections were stained with hematoxylinand eosin to differentiate cells from the surrounding collagen matrix.Large, cell-lined lumens with smaller branching lumen containing sproutswere observed USWF-exposed (FIG. 13B) samples compared to sham-exposedgels (FIG. 13A) (scale bar, 100 μm).

To assess extracellular matrix remodeling in USWF- and sham-exposedgels, collagen type-I fibers were visualized using second harmonicgeneration microscopy imaging on a two-photon microscope (FIGS. 14A-14B;scale bar, 15 microns). Human umbilical vein endothelial cells werevisualized simultaneously using intrinsic auto-fluorescence (FIGS.14A-14B). Collagen fibers were organized in linear, parallel arraysextending outwards from the endothelial cell bands in USWF-exposed (FIG.14B), but not sham-exposed (FIG. 14A) samples.

Although preferred embodiments have been depicted and described indetail herein, it will be apparent to those skilled in the relevant artthat various modifications, additions, substitutions, and the like canbe made without departing from the spirit of the invention and these aretherefore considered to be within the scope of the invention as definedin the claims which follow.

What is claimed:
 1. A vascularized engineered tissue construct, whereinsaid vascularized engineered tissue construct has a thickness inthree-dimensions of at least 2 mm.
 2. The vascularized engineered tissueconstruct of claim 1, wherein the vascularized engineered tissueconstruct is selected from the group consisting of a muscular construct,a vascular construct, an esophageal construct, an intestinal construct,a rectal construct, an ureteral construct, a cartilaginous construct, acardiac construct, a liver construct, a bladder construct, a kidneyconstruct, a pancreatic construct, a skeletal construct, afilamentous/ligament construct, a lung construct, a neural construct, abone construct, and a skin construct.
 3. A method of screening a testagent, said method comprising: providing a test agent; providing thevascularized engineered tissue construct of claim 1; contacting the testagent with the vascularized engineered tissue construct; and assayingone or more biological endpoints in the vascularized engineered tissueconstruct.
 4. The method according to claim 3, wherein the one or morebiological endpoints are selected from the group consisting ofcarcinogenicity, cell death, cell proliferation, gene expression,protein expression, cellular metabolism, and any combination thereof. 5.A method of treating a subject in need of a tissue repair or tissuereplacement, said method comprising: selecting a subject in need oftissue repair or tissue replacement; providing the vascularized tissueengineered construct of claim 1; and implanting the vascularizedengineered tissue construct into the selected subject.